The role of condensin in chromosome resolution - UCL ...

The role of condensin in chromosome resolution

Adrian Aymard Charbin

University College London

and

Cancer Research UK London Research Institute

PhD Supervisor: Dr Frank Uhlmann

A thesis submitted for the degree of

Doctor of Philosophy

University College London September 2012

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Felix qui potuit rerum cognoscere causas

Happy is he who gets to know the reasons for things

Virgil, Georgics (c. 37BC)

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Publications arising from this thesis

Condensin aids sister chromatid resolut ion by promoting their

decatenation by topoisomerase II

Adrian Charbin, Céline Bouchoux, and Frank Uhlmann. Genes and Development

(under review)

Faci le synthesis of budding yeast a-factor and i ts use to synchronise

cel ls of α mating type

Nicola O’Reilly, Adrian Charbin, Lidia Lopez-Serra, and Frank Uhlmann. 2012

Yeast 29: 233-40

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Declaration

I Adrian Charbin confirm that the work presented in this thesis is my own. Where

information has been derived from other sources, I confirm that this has been

indicated in the thesis.

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Abstract

The condensin complex is a key determinant of mitotic chromosome

architecture. In addition to its role in mitotic chromosome compaction, condensin is

required for resolution of sister chromatid linkages during chromosome segregation

in anaphase. How condensin resolves sister chromatids, and the nature of the

chromosome bridges that are characteristic of cells harboring defective condensin,

have remained topics of debate. Inactivation of topoisomerase II, the main enzyme

that removes topological interlinks that persist between DNA replication products

after their synthesis, leads to similar chromosome bridges. Here, we follow the

catenation status of circular minichromosomes of three sizes during the S.

cerevisiae cell cycle. Catenanes are produced in S-phase and, in part, they are

readily resolved, aided by physical separation of sister chromatids during mitosis.

Complete resolution, however, requires the condensin complex, a dependency that

becomes more pronounced with increasing chromosome size. Condensin and

topoisomerase II directly interact and, using purified proteins, we show that

condensin stimulates DNA decatenation by topoisomerase II in vitro. Therefore, in

parallel to promoting chromosome condensation, condensin facilitates topological

resolution of sister chromatids to secure their successful segregation to daughter

cells during cell division.

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Acknowledgement

First and foremost, I would like to thank Frank for all his advice, support and

wisdom these past four years. I have been so fortunate to be apart of his research

group, working alongside world-class scientists and learning so much. It has been

a truly great experience.

My deep gratitude goes to everyone in the laboratory who have taught me

so much but also made me laugh. Thank you especially to Chris and Maria for

running the lab so well. Without your help, my project would never have gotten as

far as it did. Thank you to Céline and Sebastian for all your guidance during my

project, I have learnt much from you both and have always enjoyed working with

you. To everyone else in the lab, thank you for all the scientific discussions and

always offering an extra pair of hands during those tricky experiments!

Thanks to all my friends who have been doing PhDs alongside me. Risa and

Vanessa for all our chats in the corridor discussing the perils of graduate work. Ali,

for our morning coffees, scientific discourses and more!

I would also like to say thank you to my family for supporting me throughout

all my studies. It is thanks to your love and encouragement that I have gotten to

where I am today.

Finally I want to thank Susie, who has filled the past four years with so much

happiness, love and laughter. I am so lucky to have you by my side, helping me

follow my dreams.

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Table of Contents

Abstract ................................ ................................ ..................... 5

Acknowledgement ................................ ................................ ....... 6

Table of Contents ................................ ................................ ....... 7

Table of Figures ................................ ................................ ....... 11

Abbreviations ................................ ................................ ........... 13

Chapter 1. Introduction ................................ ............................. 15

1.1 Historical Perspective ........................................................................................ 15

1.2 The Structural Maintenance of Chromosome Protein Family ...................... 20

1.2.1 Condensin function ....................................................................................... 21 1.2.2 Eukaryotic condensin structure ..................................................................... 22 1.2.3 Prokaryotic condensin structure ................................................................... 25 1.2.4 Condensin’s in vitro biochemical activities .................................................. 25

1.3 Chromosome Condensation .............................................................................. 27

1.3.1 Condensin’s biochemical activities drive condensation ............................... 28 1.3.2 Condensin as a catalyst driving condensation? ............................................ 29 1.3.3 Condensin could behave as a DNA linker to drive condensation ................ 29 1.3.4 Does condensin act through a co-operative mechanism? ............................. 32

1.4 Chromosome Resolution ................................................................................... 33

1.4.1 Proteinaceous linkages .................................................................................. 33 1.4.2 Topological linkages ..................................................................................... 34

1.5 Topoisomerases .................................................................................................. 35

1.5.1 Types of topoisomerase ................................................................................ 35 1.5.2 Topoisomerase II mode of action ................................................................. 37 1.5.3 The physiological importance of topoisomerase II ...................................... 40

1.6 Chromosome resolution, topoisomerase II and condensin - an unresolved

relationship .................................................................................................................. 41

1.6.1 Lessons from the rDNA locus ...................................................................... 45 1.6.2 Condensin is required for complete removal of proteinaceous links ........... 48 1.6.3 Condensin directly partners with topoisomerase II to promote decatenation 49

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1.6.4 Condensin-driven reconfiguration of mitotic chromosome topology promotes decatenation .............................................................................................. 51

1.7 An introduction to protein inactivations in S. cerevisiae ............................... 52

Chapter 2. Materials & Methods ................................ ................. 56

2.1 Yeast growth and manipulation ....................................................................... 56

2.1.1 Yeast strains .................................................................................................. 56 2.1.2 Yeast strain creation, mating and tetrad dissection ...................................... 57 2.1.3 Yeast media, cultures and synchronizations ................................................. 58 2.1.4 Anchor away strains and nuclear depletion .................................................. 58 2.1.5 Yeast transformations ................................................................................... 59

2.2 General molecular biology techniques ............................................................. 59

2.3 Protein analysis techniques ............................................................................... 59

2.3.1 Protein extract preparation ............................................................................ 59 2.3.2 Chromatin pellets .......................................................................................... 60 2.3.3 SDS-PAGE electrophoresis and western blotting ........................................ 61

2.4 Mini-chromosome purification, electrophoresis and catenane

quantification .............................................................................................................. 62

2.4.1 Catenation assay minichromosomes ............................................................. 63 2.4.2 Zymolyation of yeast cells in agarose plugs and Pulse Field Gel Electrophoresis (PFGE) ............................................................................................ 63

2.5 Cell biology and microscopy ............................................................................. 64

2.5.1 Cell cycle analysis using flow cytometry ..................................................... 64 2.5.2 Bi-nucleate cell counting .............................................................................. 64 2.5.3 In situ immunofluorescence .......................................................................... 65

2.6 Protein purifications .......................................................................................... 65

2.6.1 Purification of S. cerevisiae topoisomerase II .............................................. 65 2.6.2 Purification of S. cerevisiae condensin ......................................................... 66 2.6.3 Interaction analysis between condensin and topoisomerase II ..................... 66

2.7 In vitro assay techniques ................................................................................... 67

2.7.1 kDNA decatenation assays ........................................................................... 67

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Chapter 3. Results: Catenane Behaviour and Resolution ............. 68

3.1 Development of the catenation assay ............................................................... 68

3.1.1 Optimisation of the catenation assay ............................................................ 71 3.1.2 Identifying the minichromosome’s different topological isoforms .............. 74 3.1.3 Catenation is quantifiable at each time point ................................................ 77

3.2 Spontaneous and assisted minichromosome decatenation: ........................... 81

3.2.1 Temperature’s effect on catenation and isoform distribution ....................... 88 3.2.2 The anchor away system does not effect catenation ..................................... 92

3.3 Catenation of smaller minichromosomes is observable in wild type strains 94

Chapter 4. Results: Condensin’s Influence on Catenation ............ 97

4.1 Condensin promotes completion of minichromosome decatenation ............. 97

4.2 The effect of minichromosome size ................................................................ 105

4.2.1 Visualizing a ring chromosome’s (RCIII) different isoforms .................... 108

4.3 Condensin’s decatenation role is more pronounced on RCIII .................... 111

Chapter 5. Results: Investigating Interactions ............................ 118

5.1 Do condensin and topoisomerase II directly interact? ................................. 118

5.2 Purification of topoisomerase II and condensin ............................................ 121

5.3 Condensin and topoisomerase II directly interact ........................................ 123

5.4 Condensin stimulates in vitro DNA decatenation by topo II ....................... 123

Chapter 6. Results: a-factor Synthesis and Characterization ....... 128

6.1 Synthesis of a-factor ......................................................................................... 128

6.2 Use of a-factor to synchronise cells of α mating type .................................... 128

6.3 Little shmoo formation during a-factor-induced G1 arrest ......................... 131

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Chapter 7. Discussion ................................ .............................. 133

7.1 Catenation can be observed in minichromosomes ........................................ 133

7.2 Cohesin protects catenation ............................................................................ 133

7.3 Anaphase bridges result from persistent sister chromatid catenanes ........ 134

7.4 Condensin promotes decatenation by topoisomerase II ............................... 135

7.5 Decatenation occurs in steps ........................................................................... 136

7.6 Condensin directly interacts with topoisomerase II ..................................... 137

7.7 Future perspectives .......................................................................................... 137

Reference List ................................ ................................ ......... 141

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Table of Figures

Figure 1.1 Walther Flemming's observations of chromosomes during mitosis ...... 17 Figure 1.2 Mitotic cycle of rye (Secale cereale) chromosomes .............................. 18 Figure 1.3 Schematic representation of the condensin complex ............................ 24 Figure 1.4 Condensin subunit comparisons among different organisms ............... 24 Figure 1.5 Possible mechanisms of condensation by condensin ........................... 31 Figure 1.6 Topoisomerase II’s mode of action ....................................................... 39 Figure 1.7 Schematic of colliding replication forks, and resultant catenation ......... 44 Figure 1.8 Anaphase bridges in condensin mutants .............................................. 47 Figure 1.9 The anchor away technique of nuclear depletion .................................. 54 Figure 3.1 Map of minichromosome pS14-8 .......................................................... 70 Figure 3.2 Early electrophoresis conditions showed poor isoform resolution ........ 72 Figure 3.3 Background labelling of genomic DNA .................................................. 72 Figure 3.4 Specificity comparison of different AmpR probes .................................. 73 Figure 3.5 Wild type cells display a distinct isoform patterns ................................. 76 Figure 3.6 Enzyme digests allow for isoform identification ..................................... 76 Figure 3.7 Comparison of background deduction techniques ................................ 79 Figure 3.8 Catenane quantification of wild type cell cycle ...................................... 80 Figure 3.9 Topoisomerase II inactivation results in maximum catenane formation 82 Figure 3.10 Nocodazole arrests results in catenane persistence ........................... 84 Figure 3.11 Spindle presence during arrest promotes decatenation ...................... 84 Figure 3.12 Inactivation of cohesin results in fewer observable catenanes ........... 86 Figure 3.13 Catenane quantification for different mitotic inactivations ................... 87 Figure 3.14 Effect of temperature on catenane behaviour ..................................... 89 Figure 3.15 Effect of temperature on different isoforms ......................................... 91 Figure 3.16 Anchor away and rapamycin do not affect catenation ........................ 93 Figure 3.17 Map of minichromosome pRS316 ....................................................... 95 Figure 3.18 Catenation detection and isoform identification for pRS316 ............... 96 Figure 4.1 Condensin ts mutants all display a persistent catenane phenotype ..... 98 Figure 4.2 Condensin is required for complete decatenation ............................... 100 Figure 4.3 brn1-aa triplicates and quantification .................................................. 101

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Figure 4.4 Condensin contributes to decatenation independently of chromosome

movement ............................................................................................................. 103 Figure 4.5 Decatenation is active prior to anaphase ............................................ 104 Figure 4.6 smc2-8 releases poorly from G1 arrest ............................................... 106 Figure 4.7 Condensin depletion has a smaller effect on pRS316 catenation ....... 107 Figure 4.8 Map of ring chromosome RCIII ........................................................... 110 Figure 4.9 Resolution of RCIII by Pulse Field Gel Electrophoresis (PFGE) ......... 110 Figure 4.10 RCIII wild type timecourse and isoform identification ........................ 113 Figure 4.11 RCIII Quantification profiles .............................................................. 114 Figure 4.12 Nocodazole arrest shows persistent RCIII catenanes ...................... 115 Figure 4.13 Condensin depletion phenotype closes matches nocodazole arrest 115 Figure 4.14 Nuclear depletion of Brn1 .................................................................. 116 Figure 5.1 Co-immunoprecipitation of condensin and topoisomerase II .............. 120 Figure 5.2 Topoisomerase II purification steps .................................................... 122 Figure 5.3 Condensin interacts directly with purified topoisomerase II ................ 125 Figure 5.4 Condensin promotes decatenation of kinetoplasts ............................. 126 Figure 5.5 Condensin stimulates different topoisomerase II enzymes ................. 127 Figure 6.1 Use of a-factor to synchronise cell cycle progression of α cells ......... 130 Figure 6.2 Comparison of pheromone-induced shmoo formation of a and α cells

............................................................................................................................. 132 Figure 7.1 Schematic displaying possible excision and re-ligation of endogenous

chromosome DNA to form de novo ring chromosomes in vivo ............................ 139

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Abbreviations ARS autonomous replicating origin

ATP adenosine triphosphate

APC anaphase promoting complex

BSA bovine serum albumin

bp base pairs

Cdk Cyclin-dependent kinase

ChIP chromatin immunoprecipitation

CPC Chromosome Passenger Complex

Da Dalton (kDa, kilodalton)

DAPI 4’-6’-diamidino-2-phenylindole

DMSO dimethyl sulfoxide

DNA deoxyribonucleic acid

DNAse deoxyribonuclease

DTT dithiothreitol

ECL enhanced chemiluminescence

EM electron microscopy

EDTA ethylenediamine tetra-acetic acid

FACS fluorescence activated cell sorting

FRAP fluorescence recovery after photobleaching

FEAR Cdc14 (Fourteen) Early Anaphase Release

FRET fluorescence resonance energy transfer

G1 growth phase 1

GAL1 galactose inducible promoter 1

GFP green fluorescent protein

HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid

HRP horseradish peroxidase

Ig immunoglobin

M phase metaphase

MAT mating type

MEN Mitotic Exit Network

min minute

noc nocodazole

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OD optical density

ORF open reading frame

PBS phosphate buffered saline

PEG polyethylene glycol

PFGE pulse field gel electrophoresis

PMSF phenylmethane sulfonyl fluoride

rap rapamycin

rDNA ribosomal deoxyribonucleic acid

RNAi RNA interference

rpm revolutions per minute

S phase synthesis phase

SDS sodium dodecyl sulphate

SDS-PAGE sodium dodecyl sulphate-polyacrylamide electrophoresis

SMC structural maintenance of chromosomes

TCA trichloroacetic acid

topo I topoisomerase I

topo II topoisomerase II

ts temperature sensitive

Tris 2-amino-2-hydroxymethyl-1,3-propanediol

WT wild type

Chapter 1 Introduction

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Chapter 1. Introduction

1.1 Historical Perspective

The cell is the fundamental unit of all life we know, from single-celled bacteria

through to the billions that you consist of, reading this thesis. Yet to describe cells

as mere biological building blocks would be to overlook their immense

sophistication. If there were any word in the English language whose simplicity

most belies the complexity it represents, cell would be it. Since it was first

discovered 350 years ago the cell has remained an object of great intrigue and

even today, thousands of researchers internationally continue to make new

discoveries about how cells work. One of the cell’s most important, beautiful and

poorly understood roles is that of cell division and it is this topic that this thesis

addresses.

In 1665, the English philosopher and polymath Robert Hooke first discovered

cells when examining thin slices of cork under a coarse, compound microscope.

Seeing a multitude of tiny pores, he noted that they looked like the walled

compartments a monk would inhabit in a monastery and so called these pores

‘cells’. While Robert Hooke’s simple microscope was able to see little more than

cell walls, the compound microscope would be continuously improved upon during

the 17th century. During this time, the accumulating observations of cells from

naturalists, philosophers and the curious would eventually lead, in 1824, to Henri

Dutrochet formulating one of the fundamental tenets of modern cell theory, by

declaring that “the cell is the fundamental element of organization” (Nezelof, 2003).

In the almost two hundred years that have passed since then, cell theory has

become one of the foundations of biology. The theory states that all living things

are made of cells, that cells are the basic building units of life and that old cells

dividing into two creates new cells. While this latter point has been studied in great

detail over the years, the mechanisms underlying cell division remain incompletely

understood.

The first recorded observation of the internal organisation of cells during

division came from the second half of the 19th century, when the German biologist

Walther Flemming was using aniline dyes from a nearby chemical factory to stain


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cells before examining them under the microscope. He observed a substance that

strongly absorbed basophilic dyes, which formed apart of threadlike structures

inside the cell nucleus; he would names these chromatin and chromosomes

respectively. Through studying cell division, Flemming observed and carefully

recorded (Figure 1.1) the distribution of chromosomes into the daughter cells, a

process he called mitosis after the Greek word for thread. Following this, Flemming

concluded for the first time that all cell nuclei came from another predecessor

nucleus (Paweletz, 2001). Meanwhile, at roughly the same time, the Augustinian

friar and scientist Gregor Mendel was studying heredity in pea plants, observing

how his plants inherited certain traits with particular patterns. Unbeknownst to him,

his observations would one day gain him fame as the founder of genetics. However,

Flemming was unaware of Mendel’s work, and the significance of Flemming’s

microscopy observations and their ability to describe a mechanism for inheritance

would remain unrealised until the rediscovery of Mendel’s studies almost two

decades later in the early 1900’s.

In the 20th century, it was shown that Flemming’s chromatin was comprised

of DNA and associated proteins, and that the DNA itself was the hereditary vehicle

behind the process of inheritance Mendel had observed (Morgan, 1915). In turn,

chromosomes would be fully described as organised chromatin structures, with

each chromosome containing a single piece of DNA. However, it was quickly

noticed that the presence of chromosomes was not a permanent feature in the cell.

Indeed, during interphase when the cell is not undergoing cell division, individual

chromosomes are not visible. Upon entry into mitosis, the distinct chromosome

‘threads’ could be seen forming through ‘chromosome condensation’ (Figure 1.2) a

process driving the cytological manifestation of the chromosomes themselves.

Thus, it became apparent that the appearance of distinct chromosomes was

dependent on internal factors that were only active during specific times. This was

supported by demonstrations where interphase cells were merged with mitotic cells

causing condensation of the interphase DNA (Rao and Johnson, 1970).


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Figure 1.1 Walther Flemming's observations of chromosomes during

mitosis

Hand drawn illustrations of mitotic chromosomes, from Flemming’s 1879 paper “Beitrage zur Kenntniss der Zelle und Ihrer Lebenserscheinungen”.


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Figure 1.2 Mitot ic cycle of rye (Secale cereale) chromosomes

Scanning electron microscopy (SEM) images of chromosomes undergoing condensation. In interphase (A), DNA is in an uncondensed state, such that individual chromosomes are not identifiable. When condensation begins (B), then individual chromosomes become visible. Condensation is greatest in late metaphase (E), when separation of chromatids becomes visible. After successful segregation of the sister chromatids (F), the chromosomes begin to de-condense. Images from (Zoller et al., 2004).


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The compaction of chromatin that occurs when cells enter mitosis is

probably the most iconic process of dividing cells and despite its seemingly

superficial nature (Figure 1.2), chromosomes condensation is a ubiquitous process

in eukaryotic cells. Such is the widespread prominence of condensation in the cell

cycle that observers began to suspect that process goes beyond the task of just

making chromosomes smaller. Indeed, it is currently hypothesised that

chromosome condensation is required to resolve several important structural

problems associated with proper DNA segregation in mitosis (Baxter and Aragon,

2012). Firstly and perhaps most obviously, chromosomes are longer than the

length of the cell in which they reside. A typical human cell has a diameter of 10

micrometres, while the approximate length of DNA in its largest chromosome is 2

centimetres. Indeed the total length of cellular DNA is up to a hundred thousand

times the length of the cell itself. Therefore if chromosomes are not properly

condensed, they are unlikely to segregate properly, potentially becoming entrapped

during cytokinesis (Koshland and Strunnikov, 1996a, Hirano, 2000). Secondly,

interphase chromosomes mix, particularly in transcriptionally active areas and this

needs to be separated during mitosis. Finally, after DNA replication is complete the

two newly synthesised DNA strands, or sister chromatids, are extensively linked by

both DNA intertwines, or catenanes (Sundin and Varshavsky, 1980), as well as

proteinaceous links (Michaelis et al., 1997). These topological and proteinaceous

links must be removed if the sister chromatids are to be successfully pulled apart to

opposite poles of the cell by the mitotic spindle. The removal of all topological and

proteinaceous links between sister chromatids is called chromosome resolution, a

process describing the spatial individualization of the chromatids from each other. It

has been proposed that chromosome condensation, an ordered process structuring

DNA into individual rod-shaped chromosomes during mitosis, could drive the

resolution of any catenation between chromatids as well as shortening their length,

thus allowing their proper segregation during anaphase (Hirano, 1999).

Given its importance in chromatid resolution and segregation during mitosis,

chromosome condensation can be unequivocally classified as an essential

housekeeping function, indispensible for cell proliferation. However, the driver of

condensation had not been discovered until recently. About 15 years ago parallel

studies in several laboratories found that a protein complex called condensin was

emerging as the primary, molecular driver of mitotic chromosome condensation


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(Kimura and Hirano, 1997a, Sutani et al., 1999, Strunnikov et al., 1995a). It is

condensin, with particular regard to its role in chromosome resolution, which will be

the focus of this thesis.

1.2 The Structural Maintenance of Chromosome Protein

Family

Condensin is a member of a family of proteins called the structural

maintenance of chromosome (SMC) proteins. SMC proteins have come to be

recognised as one of the most fundamental classes of proteins that regulate the

structural and functional organisation of chromosomes from bacteria to humans

(Ivanov and Nasmyth, 2005, Hirano, 2006). Given that SMC proteins arguably pre-

date histones, it is not surprising that SMC complexes like condensin have adopted

numerous roles throughout the course of evolution and have increasingly been

viewed as global organisers of the genome (Hirano, 2006).

The best known of the SMC proteins is cohesin, a protein complex containing

SMC1 and SMC3, which binds together sister chromatids from when they are

created in S-phase, until they are destroyed at the onset of anaphase through

cleavage by the protease seperase (Uhlmann, 2003). Sister chromatid cohesion

forms the basis for the pairwise alignment of chromosomes upon the mitotic spindle,

making possible the bi-orientated segregation of chromatids at anaphase (Tanaka

et al., 2000, Toyoda et al., 2002). In addition to cohesin and condensin, a third

SMC complex has been identified consisting of a SMC5/SMC6 heterodimer. This

complex is least well characterised of the three, but it has been implicated in having

a DNA damage prevention role as well as aiding DNA repair (Roy and D'Amours,

2011) In addition, it appears to be required for proper chromosome organisation

during meiosis (Farmer et al., 2011).

Overall, the SMC family has been so interesting to chromosome biologists

owing to several factors. First and foremost, the ubiquitous presence of these

proteins, highly conserved across all kingdoms of life, suggest a pivotal and

essential role for SMCs in the proliferation of cells. Genetic and cell-biology studies

have shown that SMC proteins are involved in chromosome segregation, structure,

regulation and recombinational repair, during both mitosis and meiosis. The next


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intriguing feature of SMC proteins is their unique protein architecture. While

originally supposed to be some form of chromatin motor, many lines of evidence

have come to show that SMC proteins represent a completely novel form of protein

machine, with a distinct tripartite ring structure, that function as a dynamic linker of

the genome (Hirano, 2006).

1.2.1 Condensin function

While condensin has not been studied in as much depth as cohesin,

multiple studies have shown that the condensin complex plays a crucial role in the

compaction of DNA to form structurally stable mitotic chromosomes and their

segregation in vivo (Strunnikov et al., 1995b, Hagstrom et al., 2002, Saka et al.,

1994a, Hirano, 2005b, Ono et al., 2003, Steffensen et al., 2001, Hudson et al.,

2003). The condensin complex is highly conserved among eukaryotes, and

prokaryotes have a condensin–like complex as well. Condensin mutants in all

eukaryotic model organisms, such as S. pombe, C. elegans or D. melanogaster, all

display similar phenotypes of uncondensed nuclei and the presence of anaphase

bridges (Bhat et al., 1996, Hirota et al., 2004, Saka et al., 1994a, Strunnikov et al.,

1995b). In addition, the inactivation of prokaryotic SMCs in E. coli, B. subtilis and C.

crescentus all show temperature-sensitive colony formation and an increase in the

number of anucleate cells at the permissive temperature, suggesting a deficiency in

chromosome segregation (Niki et al., 1991, Weitao et al., 2000, Moriya et al., 1998,

Jensen and Shapiro, 1999).

Condensin has also been reported to have a range of roles in cellular

functions during interphase, such as in the control of gene expression and

heterochromatin formation (Bhalla et al., 2002) or in X. laevis CENP-A assembly

(Bernad et al., 2011). Condensin plays a role in DNA replication checkpoint

signalling in fission yeast (Aono et al., 2002) and additionally condensin mutants

have defects in DNA damage repair (Akai et al., 2011).


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1.2.2 Eukaryotic condensin structure

Condensin is a 650 kilodalton pentameric complex consisting of a

heterodimer of SMC proteins as well as three additional non-SMC subunits (Figure

1.3). The first of the condensin subunits to be identified were proteins of the SMC

family (Strunnikov et al., 1995b), including Cut3 and Cut14 proteins in S. pombe,

their orthologs in X. laevis XCAP-E and XCAP-C and the Smc2 protein in S.

cerevisiae (Hirano and Mitchison, 1994, Saka et al., 1994a, Strunnikov et al.,

1995b). The SMC subunits form the core of condensin’s enzymatic abilities, being

both the DNA-binding subunits (Sutani and Yanagida, 1997a, Cheeseman et al.,

2002) and the DNA-dependent ATPases (Kimura and Hirano, 2000). They have a

unique domain organisation with two canonical nucleotide-binding motifs, known as

the Walker A and B motifs, which are respectively located at the N-terminal and C-

terminal domains. Between the two motifs are two long coiled-coil motifs that are

connected by a non-helical sequence. Biochemical and electron microscopy

studies have shown that the SMC monomer folds back on itself through antiparallel

coiled-coil interactions, thereby creating an ATP-binding ‘head’ domain at one end

and a ‘hinge’ domain at the other. The two SMC monomers can then associate with

each other at the hinge domain to form a V-shaped molecule (Melby et al., 1998,

Haering et al., 2002, Hirano and Hirano, 2002). Additionally, entire condensin and

cohesin complexes from both H. sapiens and X. laevis have been visualised using

electron microscopy. While cohesin displays an open and distinct ring conformation,

condensin’s ring structure appears much more closed with the arms emanating

from the hinge at a smaller angle (Anderson et al., 2002) It has been hypothesised

that these differences in structure could contribute to the different roles of

condensin and cohesin in vivo (Hirano, 2005a).

Meanwhile, the non-SMC subunits are thought to have a regulatory role on

the function of the condensin complex. In vitro studies using X. laevis egg extracts

have shown that the non-SMC subunits of condensin modulate its ATPase activity

(Kimura and Hirano, 2000), though an understanding of how they alter condensin

activity and binding to chromosomes remains to be resolved. Of these three

auxiliary subunits, the CAP-D2 and CAP-G contain HEAT (Huntingtin, elongation

factor 3, the A subunit of protein phosphatase 2A, TOR lipid kinase) repeats

(Neuwald and Hirano, 2000). A number of chromosomal proteins contain these


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repeats, which are thought to facilitate protein-protein interactions. The two HEAT

repeat containing subunits are recruited by the third subunit, CAP-H (Brn1 in S.

cerevisiae, Barren in D. melanogaster), which belongs to the kleisin family of

proteins (Schleiffer et al., 2003). Kleisins are believed to act as bridging protein

between the two SMC heads as supported by studies where the kleisin subunit was

found to directly bind the SMC head domains of recombinant human condensin

(Onn et al., 2007). For a schematic representation of the known condensin

complexes in different model organisms, please see Figure 1.4.

Given that cohesin exists only as one complex across all organisms it is

found in, the same was assumed for condensin for many years. However, a

genome database search using the human CAP-D2 sequence revealed a distantly

related protein, CAP-D3 (Ono et al., 2003). Immunoprecipitation of CAP-D3

revealed that it also associated with the core condensin subunits (SMC2 and

SMC4) along with two other subunits, CAP-G2 and CAP-H2, the former containing

HEAT repeats and the latter being a kleisin subunit. This new condensin complex

was termed condensin II (Ono et al., 2003, Yeong et al., 2003) and while only

higher eukaryotes have both condensin I and II complexes, the ratio of the two can

vary significantly between different species. Phylogenetic analysis has offered

some insight into how the two complexes might contribute to chromosome

organisation. The model organisms S. pombe, S. cerevisiae, A. nidulans and N.

crassa possess only condensin I and mitotic chromosome condensation is less

dramatic a process in these species when compared to metazoans. This has led to

the speculation that condensin II may allow organisms with larger chromosomes an

additional level of organization (Hirano, 2005b). It has recently been shown using

specific conditional knockouts of the two condensins that depleting either

condensin results in differing mutant phenotypes. Super-resolution microscopy

reveals that condensin I-depleted mitotic chromosomes are wider and shorter, with

a diffuse chromosome scaffold, while condensin II-depleted chromosomes retain a

more defined scaffold, with chromosomes more stretched and seemingly lacking in

axial rigidity (Green et al., 2012).


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Figure 1.3 Schematic representat ion of the condensin complex

Figure 1.4 Condensin subunit comparisons among dif ferent organisms


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1.2.3 Prokaryotic condensin structure

SMC proteins are widely conserved across the three domains of life. In

prokaryotes, the best-studied homolog is the SMC-related protein MukB that

functions as the core subunit of a bacterial condensin complex. It is found in a

subclass of γ-proteobacteria that includes E. coli, and despite its limited sequence

homology with other SMC proteins, MukB shares the five-domain structure

common to all SMC family members consisting of a hinge domain, long coiled-coil

arms, and head domains that form the ATP binding pockets (Melby et al., 1998).

MukB forms the complete condensin complex along two other proteins, MukE,

which binds the head domain of MukB via binding to MukF, the kleisin subunit that

is remarkably conserved in bacteria (Yamazoe et al., 1999, Schleiffer et al., 2003).

In B. subtilis the SMC protein also forms a condensin complex with two other

subunits, ScpA and ScpB (Mascarenhas et al., 2002), and the SMC has an intrinsic

DNA binding ability, distinguishable from cohesin by its ability to bind DNA in an

ATP-independent manner (Hirano and Hirano, 1998a).

1.2.4 Condensin’s in vitro biochemical act iv i t ies

Condensin’s ability to reshape chromosomes is both conspicuous and

essential in the cell cycle. While there exist several hypotheses for the mechanistic

basis of condensin’s in vivo actions, which are discussed in greater length in

chapter 1.3.1, several biochemical studies have revealed specific in vitro activities

that presumably underpin condensin’s in vivo functions.

Condensin is able to bind DNA independently of ATP hydrolysis (Kimura

and Hirano, 1997b, Strick et al., 2004), which contrasts with cohesin where it is

required for DNA binding (Onn et al., 2007). While purified condensin complexes

only display low ATPase activity, this is stimulated by the addition of DNA,

increasing ATP turnover rates by up to five-fold (Kimura and Hirano, 1997a, Kimura

and Hirano, 2000, Yoshimura et al., 2002). Interestingly, this stimulation is

dependent on the presence of the non-SMC subunits, both in vitro and in vivo, and

is not seen in their absence (Kimura and Hirano, 2000, Stray and Lindsley, 2003).

While all condensins possess ATPase activities, it remains unclear to what extent

this activity underpins condensin’s cellular functions.


26

In X. laevis, condensin isolated from mitotic cell extracts has the ability to

promote the positive supercoiling of plasmid DNA in the presence of topoisomerase

I (see chapter 1.5 for an introduction to topoisomerases) (Kimura and Hirano,

1997b). A similar activity was detected in a condensin fraction (specifically

condensin II) purified from C. elegans embryos (Hagstrom et al., 2002).

Visualisation by electron spectroscopic imaging suggests this supercoiling is driven

by the wrapping of DNA around the condensin complex in two gyres (Bazett-Jones

et al., 2002). Interestingly, condensin’s supercoiling activity is not constant and

phosphorylation appears to play a major role in its control. Several of condensin’s

non-SMC subunits are targets of mitotic kinases, notably Aurora B (Lavoie et al.,

2004, Takemoto et al., 2006, Lipp et al., 2007) and cyclin-dependent kinase 1

(Cdk1) (Kimura et al., 2001, Kimura et al., 1998, Sutani et al., 1999). Additionally, a

recent study in S. cerevisiae has shown that the Polo kinase Cdc5 directly

phosphorylates all three regulatory subunits of the condensin complex in vivo and

that consequently results in a hyperactivation of condensin’s supercoiling activity

(St-Pierre et al., 2009). Similarly, the MukB subunit of the E. coli SMC complex can

support the formation of supercoils in circular plasmids in the presence of

topoisomerase I. However, this reaction is ATP-independent and produces

negative supercoiling, which is the opposite sign to that generated by eukaryotic

condensin (Petrushenko et al., 2006).

A different change in DNA topology is observed when condensin

holocomplexes immunopurified from X. laevis egg extracts or isolated S. cerevisiae

Smc2/Smc4 dimers are incubated with nicked circular DNA in the presence of ATP

and topoisomerase II. Here, the condensin complex converts the nicked circular

DNA, via topoisomerase II-catalysed strand passage, into a positive three-noded

knot, also known as a trefoil (Stray and Lindsley, 2003, Kimura et al., 1999). This

indicates that condensin cannot only introduce positive supercoils into DNA, but

has the ability to organise two or more supercoils into an ordered, solenoidal form.

Intriguingly, this knotting activity may not require ATP hydrolysis, as evidenced by a

mutant Smc2/Smc4 dimer, defective in ATP hydrolysis, which displays similar if not

identical knotting activity (Stray and Lindsley, 2003). The E. coli MukB dimer, like

its eukaryotic counterpart, can also promote the formation of right-handed DNA

knots in the presence of topoisomerase II (Petrushenko et al., 2006).


27

An additional ATPase-independent activity is the promotion of single-

stranded DNA annealing by the S. pombe Smc2/Smc4 heterodimer (Sakai et al.,

2003, Sutani and Yanagida, 1997b). It has been speculated that this activity could

be required to remove ‘leftover’ interphase products from mitotic chromosomes,

such as RNA-DNA hybrids, in order to allow their correct segregation (Yanagida,

2000). The B. subtilis SMC dimer was shown to promote DNA reannealing in a

similar but ATP-stimulated manner (Hirano and Hirano, 1998b).

1.3 Chromosome Condensation

Chromosomes are composed of roughly equal masses of DNA, histones and

non-histone proteins. Depending on the cell cycle stage, chromosomes adopt

different conformations with varying levels of compaction (see Figure 1.2). Mitotic

chromosomes are the most distinct, reaching their most condensed forms just prior

to anaphase onset. However, even in interphase when chromosomes are most

‘uncondensed’, their length is already 1,000 fold shorter than their linear

counterparts would be. While the hierarchical packaging required to reach the level

of compaction seen in mitotic chromosomes remains a hotly debated area in

chromosome biology (Grigoryev and Woodcock, 2012), it is clear that condensation

of interphase chromatin into mitotic chromosomes is tightly linked with cell cycle

progression and requires condensin.

At the simplest level of packaging, DNA is organised by the histones. 147

base pairs of DNA wrap around each nucleosome, which are comprised of histones,

and each nucleosome is linked to the next by approximately 60 base pairs of linker

DNA (Laemmli et al., 1992). Unsurprisingly, histones were long-standing

candidates for the molecular engine underlying chromosome condensation.

However, when the histone fraction was extracted from mitotic chromosomes in

vitro, the remaining insoluble non-histone fraction, the so-called ‘chromosome

scaffold proteins’, appeared to maintain a structure similar to that of the intact

mitotic chromosomes (Adolphs et al., 1977). While histones do contribute to

condensation in protozoa and higher eukaryotes through phosphorylation of the

core histone tail (Roth and Allis, 1992, Patterton et al., 1998), they are not

responsible mechanically.


28

When high-salt extractions were performed on DNA to reveal what proteins

were tightly associated with it, two of the most abundant components of eukaryotic

chromatin were identified, topoisomerase II and condensin (Lewis and Laemmli,

1982, Earnshaw et al., 1985, Gasser et al., 1986). Soon after, topoisomerase II

dysfunction in S. pombe cells was shown to disrupt both chromosome

condensation and progression through anaphase (Uemura et al., 1987b, Holm et

al., 1985), implicating topoisomerase II as an engine of condensation. However,

topoisomerase II was also known to participate in other functions in chromatin,

such as replication and transcription, and topoisomerase II’s role in chromosome

condensation was shown not to be universal (Hirano and Mitchison, 1994, Lavoie

et al., 2002). Attention then focused onto condensin, a complex that when

inactivated or depleted from eukaryotic cells results in reduced chromosome

compaction (Saka et al., 1994b, Hirano and Mitchison, 1994, Strunnikov et al.,

1995b, Koshland and Strunnikov, 1996b, Hudson et al., 2003, Steffensen et al.,

2001). An overview of the mechanisms by which condensin could drive

condensation can be seen in Figure 1.5.

1.3.1 Condensin’s biochemical act iv i t ies drive condensation

In S. cerevisiae, condensin binding is consistent throughout the cell cycle,

implying that binding itself does not drive condensation, yet its activity is somehow

maximised upon entry into mitosis (Guacci et al., 1994). As discussed in chapter

1.2.4, condensin can drive the supercoiling and knotting of DNA and these

activities are modulated by cell cycle kinases. Indeed, condensin mutants that have

a reduced ability to be phosphorylated in vivo are defective in anaphase-specific

chromosome condensation (St-Pierre et al., 2009). Therefore, it would be

reasonable to propose that the reconfiguration of chromosome topology caused by

condensin’s biochemical activities underpins chromosome condensation. Recent

genome-wide mapping in S. cerevisiae has shown that condensin’s individual

binding sites have an average spacing of 420 kilobases of DNA between them

(D'Ambrosio et al., 2008b). Consequently, between the binding sites of each

condensin complex could be hundreds of nucleosomes that would introduce large

numbers of negative turns to the DNA (Lee et al., 2007). This suggests that taken


29

on its own, condensin’s biochemical activities would not be sufficient to obtain the

required level of chromosome compaction in mitosis. Further experiments are

required to test whether the density of condensin binding is greater on vertebrate

chromosomes, which could account for their stronger compaction during mitosis.

However, while condensin’s biochemical activities may provide an essential

contribution, it is more likely that condensin accomplishes condensation through a

catalytic, linkage or co-operative mechanism.

1.3.2 Condensin as a catalyst driv ing condensation?

If condensin’s biochemical activities alone are not sufficient to explain

condensation then perhaps their action acts as mid-point, catalysing further

topological changes by other chromosomal proteins such as the topoisomerases

(see chapter 1.5). A recent study reported an increase in positive supercoiling of

circular yeast mini-chromosomes preceding their segregation, and this is

dependent on the presence of both the mitotic spindle and condensin function

(Baxter et al., 2011). Most interestingly, they found that positively supercoiled mini-

chromosome dimers isolated from topoisomerase II-deficient cells arrested in

mitosis are more efficiently decatenated in vivo by recombinant topoisomerase II

than negatively supercoiled dimers. This therefore suggests that condensin-driven

changes in topology can render that DNA into a more favourable substrate for

topoisomerase II. However, it is not immediately clear how condensin’s promotion

of decatenation by topoisomerase II could drive chromosome condensation and so

far, there are no further reports of catalytic behaviour in condensin. Nevertheless,

this particular behaviour could have substantial importance during chromosome

resolution, a topic discussed in chapter 1.4.

1.3.3 Condensin could behave as a DNA l inker to drive condensation

Another alternative is that condensin could compact DNA by acting as a

molecular linker that brings different segments from within a single DNA strand.

This idea is supported by an interesting experiment where a single DNA strand

tethered to a paramagnetic bead was nano-manipulated allowing measurement of


30

its end-to-end extension and hence level of compaction (Strick et al., 2004). It was

seen that condensin could rapidly physically compact DNA in an ATP-hydrolysis

dependent manner. The compaction reaction was highly dynamic and reversible,

with a kinetics that did not correspond with condensin-driven positive supercoiling.

Rather, the behaviour is better explained through the contraction of the linear DNA

by bringing two segments physically together and looping out the intervening DNA.

Of course the proposal that one condensin complex can link two non-adjacent

segments of DNA leads to the obvious question, is there more than one DNA

binding site on the condensin complex?

Studies of eukaryotic condensin have so far only found evidence for one

binding site. Atomic force microscopy of S. pombe condensin appears to show that

DNA associates with the complex at the hinge domain of the SMC proteins

(Yoshimura et al., 2002). Furthermore, In vitro assays have shown that the

presence of DNA blocks the proteolytic cleavage of the Smc2 hinge domain (Onn

et al., 2007) and furthermore, isolated Smc2/Smc4 hinge domains cause a shift in

DNA electrophoretic mobility suggesting a direct interaction (Griese et al., 2010). In

prokaryotes on the other hand, examination of their SMC proteins identified a

positively charged patch in the structure of the dimerized head domains and well as

in the hinge domain that could interact with DNA (Shin et al., 2009). The

introduction of mutations that negate these positive patches reduce the shift seen

in the electrophoretic mobility of plasmid DNA suggesting that for prokaryotic SMCs,

the possibility remains that they can bind two different segments of DNA at their

head and hinge domains.


31

Figure 1.5 Possible mechanisms of condensation by condensin

A) Condensin’s biochemical activity could drive condensation through changes in the topological conformation of the DNA it binds to

B) Condensin acting as a linker between distant segments of DNA C) Condensin could interact with other condensin complexes to form

multimers, thus bringing together different DNA segments


32

Is this head/hinge double binding of DNA by the condensin complex the only

was it could link DNA? Despite condensin sharing a similar tripartite ring structure

as its fellow SMC complex cohesin, it was assumed that it did not share cohesin’s

ability to capture DNA strands within its ring structure (Ivanov and Nasmyth, 2005,

Anderson et al., 2002). However, using the same techniques used for cohesin, it

has now been shown that condensin rings can also encircle chromosomal DNA

(Cuylen et al., 2011). While cohesin rings may be able to freely slide along

entrapped DNA, condensin binding has been mapped to distinct sites (D'Ambrosio

et al., 2008b). If condensin entraps two DNA strands, this could be explained by the

constant association of condensin with one chromosome segment, while local

chromatin rearrangements are performed through the sliding of the entrapped

second strand. Another possibility is that condensin complexes entrap one DNA

strand, yet bring together separate DNA segments through association with other

complexes.

1.3.4 Does condensin act through a co-operative mechanism?

Irrespective of how condensin complexes bind DNA, it has long been

suspected that condensin function depends on interaction between the complexes.

While it had been assumed that the engagement of SMC head domains always

occurred in a heterotypic fashion (for example, Smc2-Smc4, but not Smc2-Smc2),

the result that two Smc1 head domains had homodimerised in a protein crystal

came as a surprise (Haering et al., 2004). While possibly an artefact of crystal

preparation, it led credence to the idea that condensin complexes may function in a

multimeric manner, with complexes associating to form co-operative condensation

machinery. This proposal was further supported by single molecule studies where

halving the protein concentration can completely eliminate condensation activity

(Strick et al., 2004). Microscopy observations of condensin see its distribution along

the inner axes of mitotic chromosomes, consistent with the notion that condensin

forms part of a chromosome ‘scaffold’ (Maeshima and Laemmli, 2003, Cabello et

al., 2001). However, how condensin complexes could multimerise remains unclear.

Certainly if condensin ‘scaffolds’ were to form, they could not be static structures as

shown by fluorescence recovery after photobleaching (FRAP) experiments, where


33

high turnover of condensin I was observed on mitotic chromosomes (Gerlich et al.,

2006b, Oliveira et al., 2007). While evidence for multimerisation of eukaryotic

condensins remains scarce, electron and atomic force microscopy has observed

prokaryotic SMC complexes forming linear or rosette-like aggregates

(Mascarenhas et al., 2005, Matoba et al., 2005).

In summary, of the possible mechanisms of condensin action discussed, it is

most likely that reality combines aspects from all of them. Current research into

condensin is revealing tantalisingly more about its mode of action, especially with

regards to chromosome resolution. While playing a distinct role from condensation,

it is becoming increasingly clear that condensin’s contribution to both processes

are inexorably linked.

1.4 Chromosome Resolution

Chromosome resolution is a critical step in mitosis if the sister chromatids are

to be successfully segregated and thus correct chromosome segregation is a pre-

requisite for preserving genome integrity. Cohesin and condensin are both required

to ensure faithful chromosome segregation and dysfunction of either can result in

mis-segregations and aneuploidies. While most aneuploidy human embryos are not

viable, aneuploidies that occur later in human life are often associated with the

development of malignant cancer. Chromosome instability, a term referring to a

high rate of loss or gain of whole or parts of chromosomes, is a characteristic of

most human cancers and is associated with their poor prognosis and drug

resistance (McGranahan et al., 2012, Mannini et al., 2012). In summary,

chromosome resolution describes a process of individualization, where all

topological and proteinaceous links between the newly replicated sister chromatids

are removed to produce distinct and spatially isolated entities.

1.4.1 Proteinaceous l inkages

There are two types of link between the sister chromatids that need to be

removed, the first being proteinaceous. These links are predominated by the SMC


34

complex cohesin, which establishes sister chromatid cohesin with a timing that is

tightly coupled with DNA replication so as to not allow replication products to drift

apart (Michaelis et al., 1997, Uhlmann and Nasmyth, 1998).

While cohesin binds to chromosomes before entry into S phase, it is only

during DNA replication that cohesion is established. This is accomplished by a

replication fork-associated acetyltransferase (Eco1 in S. cerevisiae, Eso1 in S.

pombe) that acetylates the Smc3 subunit of cohesin. This protein is essential for

the activation of cohesin and in cells lacking Eco1, cohesin bind the chromosomes,

but the physical linkages between sister chromatids are never established (Ivanov

et al., 2002). Cohesion establishment between sister chromatids is of utmost

importance in mitosis, essential for the correct organisation and bi-orientation of the

chromatids within the mitotic spindle in preparation for segregation during

anaphase. Sister chromatid cohesion is lost upon anaphase onset through

irreversible cleavage of the cohesin subunit Scc1 by the protease separase

(Uhlmann et al., 1999). In vertebrates, the resolution of cohesion has an additional

precursor step where a substantial portion of cohesin is removed from

chromosomes as they condense in prophase in a step mediated by mitotic kinases

(Losada et al., 2000, Losada et al., 2002, Sumara et al., 2000, Sumara et al., 2002).

As soon as cohesin is dissociated from the chromosomes, the class I histone

deacetylase Hos1 deacetylates the cohesin subunit Smc3 (Borges et al., 2010). As

non-acetylated Smc3 is required as a substrate for cohesion establishment in the

following G1 phase, the cycle is completed and ready for the next round of DNA

replication.

1.4.2 Topological l inkages

Topological links describe the physical intertwining of two newly synthesised

DNA strands produced during DNA replication. This occurs during DNA replication

as a consequence of the convergence of replisomes. Initially, the topological

challenge resulting from DNA replication was most obvious in prokaryotes, such E.

coli, which have circular genomes. The unwinding of the parental DNA for

replication in a closed, circular system would inevitably generate new strands of

DNA that were linked, or catenated, together. To overcome this topological


35

constraint of DNA replication, it was first proposed in the 1960s that a nick in one of

the parental DNA strands would eliminate any entanglements by allowing free

rotation of the DNA along its axis (Cairns, 1963). While eukaryotic chromosomes

are linear, it was found that these larger genomes behaved exactly like circular

DNA owing to the existence of domains with fixed ends that prevent any free

rotation of the DNA (Worcel and Burgi, 1972). From early on, predictions were

made that in order for any DNA to be segregated post-replication, the DNA would

need to be cut in several places (Tessman et al., 1957). These initial predictions

were made with endonucleases in mind, but as it would turn out, endonucleases

would not be responsible for these cleavages, but rather a new class of DNA

enzymes called topoisomerases. Specifically, topoisomerase I would serve as a

DNA swivel (Champoux and Dulbecco, 1972) while topoisomerase II would allow

the physical passage of one DNA strand through another via a transient double-

strand break (Brown et al., 1979, Gellert et al., 1976).

1.5 Topoisomerases

Topoisomerases are a very important class of cellular enzymes, required

for the survival of all organisms through their ability to alter DNA topology by

generating transient breaks in the double helix. There are two major classes of

topoisomerases, type I and type II, which are distinguishable by the number of DNA

strands that they cleave and the mechanism by which they alter the topological

properties of the genetic material (Wang et al., 2002, Champoux, 2001, Deweese

and Osheroff, 2009).

1.5.1 Types of topoisomerase

In eukaryotes, type I topoisomerases are monomeric enzymes that require

no high-energy cofactors and can be organised into two subclasses: type IA and

type IB. Type I topoisomerases alter topology by creating transient single-stranded

breaks in the DNA, followed by passage of the opposite intact strand through the

break (type IA) or by controlled rotation of the helix around the break (type IB). As a

consequence of this process, type I topoisomerases can moderate DNA


36

supercoiling, but are not able to remove knots or tangles from duplex DNA

(Leppard and Champoux, 2005). Eukaryotic type II topoisomerases function as

homodimers and require divalent metal ions and ATP for complete catalytic activity.

Topoisomerase II can interconvert different topological forms of DNA (also known

as isoforms) via a ‘double-stranded DNA passage reaction’, which can be broken

down into a series of distinct steps (Wang, 1998). Owing to this DNA passage

mechanism type II topoisomerases cannot only alter DNA supercoiling like the type

I enzymes, but also can remove DNA knots and catenation. It should be noted that

after action upon by topoisomerase II, the chemical structure of the ligated DNA is

identical to that of the original substrate. Hence, only the topological properties of

the double helix are changed by the actions of this enzyme.

Lower eukaryotes and invertebrates, such as S. cerevisiae, encode only a

single topoisomerase II (Goto and Wang, 1984), while vertebrate species encode

two closely related isoforms of the enzyme, topoisomerase IIα and topoisomerase

IIβ. While the isoforms are encoded by separate genes and differ in their protomer

molecular masses (170 and 180 kDa, respectively), they share similar

enzymological characteristics and high sequence homology. However,

topoisomerase IIα and IIβ do have differing patterns of expression and their own

distinct cellular roles. Type IIα is most similar to the type II topoisomerases see in

lower eukaryotes and is essential for proliferating cells to survive, with protein

levels increasing significantly during periods of cell growth. Furthermore, the

enzyme is regulated through the cell cycle with peak concentrations in G2 and M

phase (Heck and Earnshaw, 1986, Heck et al., 1988, Kimura et al., 1994).

Topoisomerase IIα is associated with replication forks and remains tightly bound to

chromosomes throughout mitosis. Thus, it is believed to be the isoform that

functions in the cell’s growth-related processes, such as DNA replication and

chromosome segregation (Wang et al., 2002, Christensen et al., 2002).

Topoisomerase IIβ on the other hand is dispensable on the cellular level and

cannot compensate for the loss of topoisomerase IIα in mammalian cells, implying

that these two isoforms do not play redundant roles in replicative processes

(Sakaguchi and Kikuchi, 2004). Instead, topoisomerase IIβ appears to be required

for proper neural development, most likely through involvement in the transcription

of developmentally or hormonally regulated genes (Ju et al., 2006, Yang et al.,

2000).


37

Prokaryotes encode two distinct type II topoisomerases, gyrase and

topoisomerase IV. Gyrase is unique in that it is the only known topoisomerase that

can actively negatively supercoil DNA and it plays an important role in regulating

the superhelical density of bacterial DNA (Levine et al., 1998). Meanwhile,

topoisomerase IV has a similar role to eukaryotic topoisomerase II, required for

decatenation of sister chromatids after DNA replication and general removal of

knots from the genome (Deweese and Osheroff, 2009).

1.5.2 Topoisomerase II mode of act ion

The ability of topoisomerase II to cleave and ligate DNA is central to all of its

catalytic functions. To perform a strand passage reaction, topoisomerase II

interacts with two DNA strands introducing a double strand break in one DNA

strand, termed the gate or G segment, and then passing a second strand termed

the T segment through the break (Figure 1.6). In the presence of Mg2+, the enzyme

can cleave the DNA by nucleophilic attack, forming a phosphotyrosine linkage

between each single strand and a tyrosine in each subunit. The covalent enzyme-

DNA linkage plays two critical roles in the topoisomerase II reaction mechanism.

Firstly, it conserves the bond energy of the sugar-phosphate DNA backbone.

Secondly, the covalent bond does not allow the cleaved DNA chain to dissociate

from the enzyme, thus maintaining the integrity of the genetic material during the

cleavage event (Wang, 1998). This is crucially important as double-stranded

breaks in DNA can have mutagenic and even lethal consequences for cells.

The next step is ATP binding, which causes the enzyme to form a closed

clamp. The closed clamp can also capture another strand (the T strand) that will

then pass through the break made in the G strand. After passing through the break

in the G strand, the T strand exits the enzyme through its carboxy terminus. ATP

hydrolysis occurs at two steps in the reaction cycle, the first ATP hydrolysed likely

assists in the passage of the T segment. The second hydrolysis step allows the

clamp to reopen, releasing the G segment, although the enzyme may initiate

another catalytic cycle without dissociating from the G strand (Harkins and Lindsley,

1998).


38

While it acts globally across all DNA, topoisomerase II appears to cleave at

preferred sites. However, the consensus sequence for cleavage is weak, and many

sites of action do not conform to it (Capranico and Binaschi, 1998). Currently, the

mechanism by which topoisomerase II selects DNA sites to act upon is not

apparent, and it is currently not possible to predict de novo whether a given DNA

sequence will support scission (Deweese et al., 2008). Most likely, the specificity of

topoisomerase II-mediated cleavage is determined by a combination of the local

flexibility, structure or malleability of the DNA that accompanies the sequence, as

opposed to the direct recognition of the bases that comprise that sequence (Velez-

Cruz et al., 2005).


39

Figure 1.6 Topoisomerase II ’s mode of act ion

Type II topoisomerases bind two separate segments of DNA and then create a double-stranded break in one of the segments. After translocating the T segment through the cleaved nucleic acid ‘gate’ (G segment), the enzyme ligates the cleaved DNA, releases the translocated segment through a gate in the protein. Finally, the protein gate is closed and the enzyme regains the ability to start a new round of catalysis (adapted from Deweese et al., 2008).


40

1.5.3 The physiological importance of topoisomerase II

The correlation between faithful segregation of sister chromatids in mitosis

and cancer cell development was touched upon earlier in this introduction. SMC

proteins are required for proper chromosome segregation and their dysfunction can

lead to the mis-segregations and aneuploidies associated with malignant tumours.

However, the unique and critical role topoisomerase II has in managing DNA

topology makes it perhaps the most important enzyme to understand with regards

to chromosome segregation. Indeed, topoisomerase II has become a prime target

for manipulation by anti-cancer drugs (Fortune and Osheroff, 2000, McClendon and

Osheroff, 2007).

Unlike most other protein-targeted drugs, which kill cells by robbing them of

an essential enzyme activity, topoisomerase-targeted drugs exploit the potentially

lethal nature of topoisomerases. During the strand passage reaction, the

topoisomerase II-DNA cleavage complexes are normally short-lived and readily

reversible, with the DNA cleavage/ligation equilibrium of the enzyme greatly

favouring ligation (Bender et al., 2008, Mueller-Planitz and Herschlag, 2008).

Therefore, topoisomerase-targeted drugs ‘poison’ topoisomerase II by increasing

the steady-state levels of the DNA cleavage complexes (Fortune and Osheroff,

2000, McClendon and Osheroff, 2007). This action converts topoisomerases into

potent physiological toxins, resulting in the generation of DNA double strand breaks.

Subsequently, the resultant mass of DNA damage can overwhelm the cell’s repair

pathways resulting in apoptosis (Roos and Kaina, 2012).

These drugs have proven to be powerful tools for oncologists, but growing

evidence now suggests that topoisomerase II-mediated DNA cleavage can trigger

chromosomal translocations that lead to the development of specific types of

leukaemia. It has been shown that up to 3% of patients who take topoisomerase II-

targeted drugs eventually develop acute myeloid leukaemia (Felix et al., 1995, Felix,

1998, Bender et al., 2008). Therefore, given topoisomerase II’s role in both healthy

cell division and the development of malignancies, gaining a better understanding

of this enzyme’s function in chromosome resolution and its relationship to other

chromosomal proteins like condensin, is of great importance.


41

1.6 Chromosome resolution, topoisomerase II and

condensin - an unresolved relationship

From bacteria to humans, DNA is globally underwound, or negatively

supercoiled, by approximately 6% (Bauer et al., 1980). This state is important as

duplex DNA is merely the storage form of the genetic information and in order to

replicate or express this information, the two strands of DNA must be separated. As

global negative supercoiling of the genome imparts increased single-stranded

character to the double helix, it therefore also facilitates strand separation (Espeli

and Marians, 2004). While negative supercoiling may promote many nucleic acid

processes, DNA overwinding, or positive supercoiling, inhibits them. During

replication the linear movement of DNA enzymes, such as the helicases and

polymerases of the replisome, compresses the turns of the double helix into a

shorter region ahead of the replication fork causing it to become increasingly

overwound. The positive supercoiling that results makes it progressively more

difficult to open the two strands of the double helix and eventually this can block

essential nucleic acid processes (Travers and Muskhelishvili, 2007). Normally, type

I topoisomerases are able to relieve this supercoiling but in the latter stages of

replication when replisomes approach each other, the two replication forks impinge

on each other and there is no longer room for a type I topoisomerase to relax the

positive supercoiling (Murray and Szostak, 1985, Sundin and Varshavsky, 1980,

Sundin and Varshavsky, 1981). By allowing the replisomes to rotate along the axis

of the DNA strand, the torsion building up ahead of the fork is relieved. A

consequence of this is that pre-catenated DNA molecules are created behind the

fork, that upon replication completion, result in fully catenated DNA daughter

molecules (Figure 1.7) (Zechiedrich and Cozzarelli, 1995, Lucas et al., 2001).

These topological links between the sister chromatids must be removed should

they be successfully segregated during anaphase and this is performed by

topoisomerase II, the only enzyme capable of passing one DNA strand through

another via a transient double strand break (Wang, 2002). Therefore,

topoisomerase I and II mediated topological transitions at the replication forks

ensure proper fork progression and stability and prevent activation of the DNA

damage checkpoint (Bermejo et al., 2007). Topoisomerase II is therefore essential

in mitosis and mutational analysis from various model organisms have shown it to


42

be the main decatenation driver, with mutants having distinct phenotype

characteristics including poorly condensed chromosomes as well as non or mis-

segregation of the sister chromatids in anaphase (Downes et al., 1991, Holm et al.,

1985, Uemura et al., 1987a).

While the importance of topoisomerase II in decatenation was indisputable,

several nagging issues persisted that questioned whether this enzyme could

actually represent the whole story. The most pertinent question was how the cell

could be sure its whole genome had been fully disentangled by anaphase onset?

While topoisomerase II is an exceptional enzyme in its capabilities, its main

shortcoming is that its actions are bi-directional. Not only can it remove topological

linkages, but it can introduce them as well. In most circumstances, when two

entangled DNA strands are decatenated by topoisomerase II, the liberated strands

are then likely to move apart spatially. Not only are the two strands released from

opposing sides of the topoisomerase enzyme, but their further separation is also

energetically favourable through Brownian motion. Therefore, the likelihood that

two strands come close enough together again to be recatenated is low. However,

in a situation where the two strands are physically constrained, such as if a cohesin

complex entraps both, then the probability of recatenation is greatly increased.

Various arguments have been made to overcome this issue, the first being that

since topoisomerases are ATPases, they are capable of catalysing an end point

that is beyond a thermodynamically stable equilibrium (Vologodskii et al., 2001). In

addition, it has been shown that topoisomerase II can detect specific DNA topology

as suggested by their ability to interact with DNA crossovers (Dong and Berger,

2007, Zechiedrich and Cozzarelli, 1995). However, these arguments are not fully

satisfactory in explaining how the complete decatenation of the sister chromatids is

ensured.

The idea that condensin may contribute to resolution of sister chromatids in

addition to its role in chromosome condensation was reinforced with the

characterization of the complex in S. cerevisiae (Freeman et al., 2000). When

compared to vertebrates, the chromosomes of budding yeast do not attain the

same high levels of condensation in mitosis. In fact in S. cerevisiae, condensin-

driven condensation causes only a 1.5-fold compaction of the chromosome arms

and for successful chromosome segregation, only the longest chromosomes

require this shortening (Guacci et al., 1994). Yet in the absence of condensin, all


43

the chromosomes fail to segregate (Bhat et al., 1996). This situation makes the

sister chromatid sorting function of condensin even more apparent. Recently,

studies of S. cerevisiae rDNA have been key in developing our understanding of

condensin’s resolution function.


44

A

B

Figure 1.7 Schematic of col l id ing repl icat ion forks, and resultant

catenation

A. As replication forks approach each other head on, helicases are no longer able to relieve the building superhelical tension in the DNA ahead of the fork. Through rotation of the replisomes this tension is relieved, but topological linkages, or catenation, is established between the sister chromatids.

B. A schematic of a topological link between DNA strands belonging to different sister chromatids. This link must be resolved to allow proper DNA segregation or else risk DNA damage.


45

1.6.1 Lessons from the rDNA locus

The budding yeast rDNA locus consists of between one and two hundred

9.1kb DNA repeats located on the longest arm of chromosome XII. At this site,

ribosome biogenesis occurs as the nucleolus assembles on top of the locus. It has

been previously noted that sister chromatids remain connected at the rDNA until

mid-anaphase independently of the cohesin complex and that to remove this

connection, both topoisomerase II and condensin activities are required (D'Amours

et al., 2004, Sullivan et al., 2004). The phosphatase Cdc14, one of the most

important down regulators of mitotic cyclins during mitotic exit in S. cerevisiae,

appears to activate resolution of the rDNA locus, though a mechanistic explanation

of this remains undetermined. The rDNA locus was interesting to study, as not only

was it easy to visualise using immunofluorescence microscopy, but the condensin-

mediated compaction of the locus could be functionally separated from condensin-

mediated resolution. This is because compaction, but not resolution, depends on

Aurora B kinase. Upon inactivation of the kinase, uncondensed rDNA sister

chromatids succeeded in full rDNA disjunction (D'Amours et al., 2004, Sullivan et

al., 2004). However, despite these uncondensed sister rDNA loci being fully

resolved, they fail to be efficiently segregated in the dividing cell (Sullivan et al.,

2004). Overall, this suggests that the most important function of condensin in rDNA

resolution is not chromosome compaction, but resolution.

Examination of this situation is made even more interesting by considering

the role rDNA plays within the cell. As the site of ribosome biogenesis, the rDNA

locus is area of intense and continual transcription by RNA pol I and RNA pol III. It

is well known that specific DNA structures are formed during the process of

transcription and these are recognised and relaxed by topoisomerases (Mondal

and Parvin, 2001, Osborne and Guarente, 1988). Besides this, in S. cerevisiae

transcription continues non-stop through the cell cycle unlike in higher eukaryotes

where it pauses during anaphase (Elliott and McLaughlin, 1979, Sullivan et al.,

2004). Given that rDNA transcription generates a large number of DNA structural

forms, it is conceivable that this locus would impede its own topological resolution

without the concerted action of condensin and topoisomerase II. Correspondingly, it

has been shown that growing S. cerevisiae cells in the presence of a transcription-

inhibiting drug enhances rDNA segregation (Tomson et al., 2006). Given that the


46

requirement for condensin in rDNA partitioning and its massive enrichment at this

loci (Freeman et al., 2000), it was therefore feasible that condensin’s specific role

was to aid resolution in topologically complex areas of high DNA transcription.

However, a study then demonstrated that cell cycle down-regulation of rDNA

transcription inversely correlated with condensin’s binding efficiency to the rDNA

repeats (Wang et al., 2006). Taken together, these studies painted a conflicting

picture of what was happening. On the one hand, it seemed that high levels of

transcription were conflicting with resolution, perhaps by preventing stable binding

of condensin to the DNA. On the other, while transcription continues unabated at

the rDNA loci, wild type cells can correctly segregate chromosomes during

anaphase. This specific situation was finally clarified in a study where it was shown

that the sister-rDNA segregation defect seen in condensin mutants could be

overcome by ectopic expression of a foreign topoisomerase II (D'Ambrosio et al.,

2008a). This result implied that it was indeed catenation preventing sister-rDNA

segregation but that the endogenous S. cerevisiae topoisomerase II was ineffective

in decatenating the locus without condensin.

Apart from S. cerevisiae, depletion of condensin in a range other cells

resulted in phenotypes that closely resembled those seen in topoisomerase II

inactivations (Strunnikov et al., 1995b, Bhat et al., 1996, Hagstrom et al., 2002,

Hudson et al., 2003). These include poorly condensed and mis-segregated mitotic

chromosomes, but the most striking feature are anaphase bridges (Figure 1.8).

These result from the mitotic spindle pulling incompletely-resolved sister

chromatids apart to opposing poles of the cell and the unresolved DNA is seen

stretched as a bridge between the two spindle poles (Lavoie et al., 2002, Chan et

al., 2007). The exact cause of these bridges has not been determined and their

presence in condensin mutants has been attributed to many different causes, from

the abnormal compaction of DNA in early mitosis (Hirano, 2005a) to the premature

loss of compaction in early anaphase (Gerlich et al., 2006a, Vagnarelli et al., 2006).

Conversely, the similarity between condensin and topoisomerase II inactivation

phenotypes has led to the proposal that condensin may promote DNA decatenation

through interaction with topoisomerase II (Bhat et al., 1996, Coelho et al., 2003,

Sullivan et al., 2004). Despite the diverse range of proposals for condensin’s role in

chromosome segregation, the most likely mechanisms can be summarised in three

different scenarios (Cuylen et al., 2011), discussed in the following three chapters.


47

Figure 1.8 Anaphase bridges in condensin mutants

Immunofluorescence microscopy images of anaphase bridges in DAPI-stained DNA presenting in (top) S. cerevisiae condensin mutants (Lavoie et al., 2002), and in (bottom) H. sapiens condensin mutants (Chan et al., 2007).


48

1.6.2 Condensin is required for complete removal of proteinaceous

l inks

As introduced before, the main proteinaceous linker connecting the sister

chromatids prior to anaphase is cohesin (see chapter 1.4.1). Therefore it is

possible that depletion of condensin results in the incomplete removal of cohesin

and this has been demonstrated in metazoan cells. In nocodazole-arrested HeLa

cells depleted of condensin I (but not condensin II) still retain small amounts of

cohesin on the chromosome arms (Hirota et al., 2004), impairing the resolution of

chromosome arms normally observed under those arrest conditions, suggesting

that condensin is required for complete removal. In S. cerevisiae condensin

mutants, there is also an apparent failure to remove all cohesin from chromosome

arms during mitosis (Renshaw et al., 2010) and even in meiosis, condensin

mutants display telomeric segregation defects that can be reduced by over

expression of seperase (Yu and Koshland, 2005). The kleisin subunit of cohesin is

the cleavage target of separase and it has been shown that kleisin phosphorylation

by PLK1 renders it a better substrate (Alexandru et al., 2001). Correspondingly, a

reduction was observed in condensin mutants in the localization of PLK1 and the

phosphorylation of cohesin’s kleisin subunit during the first meiotic division (Yu and

Koshland, 2005). However it is not clear if condensin could recruit PLK directly to

cohesin, as while both SMC complexes are loaded onto the DNA by a shared

loader complex (Scc2/Scc4) (Ocampo-Hafalla and Uhlmann, 2011), their final

localization on the chromosome arms appear separate (D'Ambrosio et al., 2008b).

Instead of a direct role, condensin could destabilise cohesin binding through the

manipulation of mitotic chromosome structure. In support of this, the anaphase

movement dynamics of fluorescently labelled S. cerevisiae chromosomes was

tracked. Upon anaphase onset, segregation of the sister chromatids begins at the

centromeres before moving along the chromosome arms with delays that increase

towards the telomere. By promoting additional degradation of the kleisin at

anaphase onset, and thus increasing cohesin removal, removes the observed

delays. This implies that an uncleaved population of cohesin underlies the

sequential stretching of DNA along the chromosomes arm. When this residual

cohesin is cleaved by seperase, the chromosome arms spring back. However, in

condensin mutants this elastic action is not seen. This leads to the proposal, which


49

has been supported by mathematical modelling, that condensin-dependent

manipulation of DNA topology promotes removal of these residual uncleaved

cohesins (Renshaw et al., 2010). Countering this, are studies done in human cells

where depletion of Wapl results in much higher levels of cohesin being bound to

the chromosomes. Despite the much higher levels of cohesin present, no

segregation problems are observed (Shintomi and Hirano, 2009). As an interesting

aside, there have also been reports that topoisomerase II functions in the cohesin

cycle, hinting at the possible of a broad relationship between the enzyme and the

SMC family (Tapia-Alveal et al., 2010).

1.6.3 Condensin direct ly partners with topoisomerase II to promote

decatenation

DNA replication results in the topological linkage of the sister chromatids

and this catenation is removed by topoisomerase II prior to anaphase onset.

However, topoisomerase II is a bi-directional enzyme and as such the probability of

these enzymes disentangling entire chromosomes on their own is very unlikely. As

such, condensin has been proposed to promote of complete decatenation via a

direct interaction with topoisomerase II. In prokaryotes, this proposal has recently

received significant credit on the back of two studies demonstrating that the E. coli

SMC protein MukB directly binds to and stimulates the activity of topoisomerase IV

(Li et al., 2010, Hayama and Marians, 2010). In vitro assays demonstrated that the

incubation of DNA with MukB prior to the addition of topoisomerase IV promoted

the relaxation, and to a lesser extent decatenation, abilities of the enzyme. The site

of interaction between the proteins was mapped via mutations to the MukB hinge

domain and the C-terminal domain of the topoisomerase IV ParC subunit. This

stimulatory effect could be a consequence of MukB binding simply boosting

enzyme activity. Alternatively, MukB could preferentially bind sites of DNA

catenation and then recruit topoisomerase IV to them.

Given the interaction observed in prokaryotes, it would not be unreasonable

to predict a similar relationship in eukaryotes. Unfortunately, establishing where

condensin directly interacts with topoisomerase II has been difficult and produced

conflicting reports. The idea was first floated by the findings that mutations in the S.

pombe genes encoding the Smc4 subunit and topoisomerase II are synthetic lethal


50

(Saka et al., 1994b). This was then followed by reports that the D. melanogaster

kleisin subunit Barren co-localised with topoisomerase II and activated the

enzyme’s activity in in vitro assays (Bhat et al., 1996). However, while later studies

confirmed that D. melanogaster cell extracts depleted of Smc4 lose their

decatenation abilities, no direct interaction between condensin and topoisomerase

II was observed (Coelho et al., 2003) and no direct interactions were observed in S.

cerevisiae either (Bhalla et al., 2002). To further muddy the waters, extracts from

mitotic X. laevis cells depleted of condensin showed no reduction in decatenation

ability (Cuvier and Hirano, 2003). As such, there remains no overall consensus in

eukaryotes on whether condensin direct binds topoisomerase II or not, with

variation observed both between and within model organisms.

However, condensin could still directly promote decatenation through the

recruitment of topoisomerase II to catenation sites on sister chromatids. This idea

could explain the reduction in topoisomerase II staining on mitotic chromosomes

spreads seen in S. cerevisiae condensin mutants (Bhalla et al., 2002). In vitro

experiments show that condensins can stimulate plasmid knotting and

preferentially bind structured DNA substrates, which hint at a possible affinity for

DNA crossover sites (Kimura and Hirano, 1997b, Sakai et al., 2003). The

enrichment of condensin in the topologically complex DNA landscape of rDNA loci

in S. cerevisiae also lends credence to this line of thought (Freeman et al., 2000,

Wang et al., 2005, D'Ambrosio et al., 2008b). However, the prospect of condensin

acting as a recruitment agent for topoisomerase II remains unconvincing. Firstly,

under some conditions the requirement for topoisomerase II function is no longer

needed during rDNA segregation in anaphase, while condensin remains essential

(D'Amours et al., 2004). While CHIP on CHIP mapping reveal a loose co-

localization along the chromosomes (D'Ambrosio et al., 2008b), this is conflicted by

immunofluorescence studies of mitotic chromosomes (Maeshima and Laemmli,

2003). Additionally, overexpression of topo II rescues co-localization in condensin

mutants, but not function (Bhalla et al., 2002). Finally, the depletion of condensin

subunits in metazoan cells does not significantly affect chromosomal

topoisomerase II levels (Hudson et al., 2003, Hirota et al., 2004).


51

1.6.4 Condensin-driven reconfiguration of mitot ic chromosome

topology promotes decatenation

If condensin does not directly interact with topoisomerase II, it could still

direct its behaviour. While condensin does not possess topoisomerase’s ability to

cut and translocate DNA strands, it does have its own biochemical activities that

alter DNA topology as introduced in chapter 1.3.1. Condensin’s reconfiguration of

chromosome topology during chromosome condensation could promote

decatenation and proper resolution on both local and global levels.

On a local level, it has been shown that positively supercoiled mini-

chromosome dimers are more efficiently decatenated by topoisomerase II than

negatively supercoiled dimers (Baxter et al., 2011). Given that the positive

supercoiling of these mini-chromosomes is dependent on the presence of

condensin (as well as the mitotic spindle), it suggests that condensin’s supercoiling

activity could generate DNA substrates, which are more amenable to decatenation,

thereby pushing topoisomerase II’s reaction equilibrium towards complete

resolution. On a global level, the compacting activity of condensin within (but not

between) mitotic chromosomes means that catenated sister chromatid DNAs are

being pulled away from each other in a tug-of-war manner. Thus, it is not only

energetically favourable for topoisomerase II to separate the DNAs, but upon

decatenation, the sister chromatid DNAs are immediately physically isolated

making any recatenation impossible.

Of course, condensin-driven condensation could also promote the types of

resolution already discussed. For example, the stiffening of the chromatid fibre

caused chromosome condensation could allow the transmission of mechanical

forces generated at the centromeres by the pulling of the spindle out to the

chromosome arms, thereby tearing apart any residual cohesin linkages that had

not been cleaved by separase (Renshaw et al., 2010).

Condensin is required in three major mitotic events: chromosome

condensation, chromosome resolution of proteinaceous linkages and chromosome

resolution of topological linkages. It has become increasingly apparent that all three

processes are intimately linked, with condensin being the common denominator.

However, the relative contribution of condensin, particularly in regards to


52

chromosome resolution, as well the mechanism of its action remain unclear. This

thesis presents work that aims to address these shortcomings in our knowledge of

condensin.

1.7 An introduction to protein inactivations in S. cerevisiae

In the lab, we possess a range of temperature sensitive (ts) mutants suitable

for use in functional studies and to date, the majority of what is known about yeast

condensin has been learnt from the use of these ts alleles. In a ts mutant, a specific

protein is altered such that at the permissive temperature, usually 25°C, the protein

behaves as if wild type. However, upon temperature shift to the non-permissive

temperature, usually 37°C, then the mutant protein will cease to function and be

inactivated. At the start of our project, we used ts mutants extensively to inactivate

proteins of interest and examine the consequence on catenation. Given the sizable

collection of ts mutants already in possession by the laboratory, we did not need to

create any further ts mutants in order to examine all the proteins we were

interested in.

However, the use of ts mutants does present some problems. Firstly, some

ts mutants have a leaky response to temperature shift, resulting in only a partial

inactivation of the target protein. Secondly, not all the ts mutants are fully

characterised, such that while we know the protein is inactivated at the non-

permissive temperature, the exact mechanism of its inactivation is unclear. For

example, if we shift a condensin ts allele, such as smc2-8, to a non-permissive

temperature we are unsure of the exact effect on the protein. Does the shift cause

the whole condensin pentameric complex to disassemble or does the complex

remain intact but its functional activities are stopped? Does the protein remain

bound to DNA and if so, could it still have some persistent effect on DNA topology,

perhaps through interaction with another chromosomal protein? This lack of

knowledge on the exact nature of the inactivation means that when we started

contemplating the mechanisms behind our results, we were unable to have much

confidence in the models we proposed.

Separate to this uncertainty about the inactivation, the temperature shift

required to inactivate the target protein, usually a change from 25°C to 37°C, will of


53

course itself have an effect on the cells (Morano et al., 2012). At the higher

temperature, the cell’s stress response will be activated and when considered in

the light of our catenation assay, we needed to be sure that the sometimes-subtle

changes observed in our data were a result of a specific target protein inactivation

and not the cell’s innate stress response. While we performed the necessary

control experiments (see chapter 3.2.2 of the results) at the start of our project,

these limitations were deemed acceptable because the only other alternative for

inactivating target proteins was the Degron method (Dohmen et al., 1994), but this

too uses a temperature shift to initiate degradation.

However, shortly after starting this PhD project, a paper was published

detailing a novel technique for the specific depletion of a protein of interest from the

nucleus of S. cerevisiae called anchor away (Haruki et al., 2008b). The concept

was simple: to sequester, in a ligand-dependent manner, a gene product of interest

from its functional compartment to a different one, where it is anchored to a suitable

protein receptor (the anchor) and cannot exert its function. Thus, a target protein

could be specifically depleted from the nucleus by being transported out of the

nucleus and tethered to a receptor protein located in the cytoplasm. This was

achieved through the use of the human 12kDa, FK506 binding protein (FKBP12)

and the 11kDa, FKBP12-rapamycin-binding (FRB) domain of human mTOR, which

were fused to the anchor and target proteins, respectively. Rapamycin binds to the

FKBP12 domain where it forms an interaction surface for the FRB domain to

establish a tight tertiary complex of nanomolar dissociation constant (Chen et al.,

1995). By taking advantage of the massive flow of ribosomal proteins through the

nucleus during maturation, when rapamycin is added and there is formation of a

tertiary complex composed of the anchor, rapamycin, and the target, then this

results in the rapid depletion of the target from the nucleus (Figure 1.9).

The new anchor away technique was exciting as it provided a new means

by which to inactivate a particular protein, in this case through the specific depletion

of the target protein from the nucleus. Given the problems arising from use of ts

alleles, using anchor away offered many advantages, the most important being that

any temperature shifts could be avoided and all experiments performed at 25°C.

Regarding the efficiency of the depletion, by probing Western blot samples of

chromatin pellets, we observed an approximately 80% depletion of the target

protein thirty minutes after rapamycin addition.


54

Figure 1.9 The anchor away technique of nuclear deplet ion

This simplified schematic shows how the addition of rapamycin results in the formation of a ternary complex including the target protein with the epitope tag FRB and the (non-nuclear) anchor-FKBP12. The end result is the depletion of the target protein from the cell’s nucleus.


55

Therefore, when presenting our results we have often done the inactivation

of the protein using both ts alleles and the anchor away technique and this is

because of several reasons. Firstly, by the time we had obtained the anchor away

strain in the lab we had already done many of experiments using the ts alleles.

Later on in the PhD, it would often be faster to do experiments using the ts strains

as it could take time to correctly tag or cross strains for use with anchor away.

Furthermore, and perhaps most importantly, by repeating key experiments using

both approaches we were able to really ensure that the resulting phenotypes

observed were indeed a consequence of inactivating that specific protein and not a

side effect of temperature, strain sickness or incomplete inactivation of the target

protein. As such, it acted as a control and confirmation of our observations and

thus both approaches were used throughout.

Use of the anchor away strain also resulted in a small side project. In

research labs using S. cerevisiae as their model organism, the ability to

synchronise cell populations using the mating pheromone α-factor has proven

invaluable, especially for cell cycle studies. Given its common use, the α-factor

response pathway has also become an important model to study the molecular

mechanism of G-protein coupled receptor signalling (Vallier et al., 2002). However,

only cells of the a mating type will respond to this pheromone. As it happened, the

anchor away cell lines were of the α mating type, and as such do not respond to α-

factor but rather to a-factor, a farnesylated and C-terminally methylated 12 amino

acid peptide. Because of its more difficult chemical synthesis, a-factor is not readily

available and consequently the a-factor response in S. cerevisiae is poorly

characterised.

Given our extensive use of anchor away strains throughout this project, and

resultant high consumption of a-factor, we collaborated with the peptide synthesis

laboratory in our institute. While they developed a new and improved strategy for a-

factor production based on solid-phase peptide synthesis, we characterised the

successful use of the resultant a-factor in synchronization of cell populations

(O'Reilly et al., 2012). Results from this characterization of a-factor and

comparisons with α-factor can be seen in chapter 6 of the results.

Chapter 2 Materials and Methods

56

Chapter 2. Materials & Methods

2.1 Yeast growth and manipulation

2.1.1 Yeast strains

The strains used in this study were of the W303 background, or were

backcrossed against this background, with the exception of the brn1-9 (Y3939) and

smc4-1 (Y3954) strains that were of S288c background. Strain details are listed

below:

Strain

Number

Genotype

K699 MATa , ade2-1, trp1-1, can1-100, KAN, his3-11, ura3-52, GAL,

psi+, (w303 wild type)

Y216 MATα, SCC1- HA6::HIS3

Y390 MATa ATa, BRN1-HA6::HIS3

Y2665 MATa , w303 wild type containing pS14-8(LEU2)

Y3277 MATa , TOP2-PK3::LEU2

Y3278 MATa , BRN1-HA6::HIS3, TOP2-PK3::LEU2

Y3939 MATa , brn1-9, pS14-8(LEU2) (S288c background)

Y3940 MATa , NET1-GFP::TRP1, ycg1-10, pS14-8(LEU2)

Y3954 MATa , smc4-1, pS14-8(LEU2) (S288c background)

Y3976 Topo II purification strain. See Worland & Wang 1989

Y3993 Condensin purification strain. See St-Pierre et al. 2009

Y4007 MATa , top2-4, pS14-8(LEU2)

Y4029 MATa , scc1-73, pS14-8(LEU2)

Y4031 MATa , SCC1-HA6::HIS3, TOP2-PK3::LEU2

Y4113 MATα, tor1-1, fpr1::NAT, RPL13A-2xFKB12::TRP1, smc2-

FRB::HIS3, MET3pr-HA-CDC20::URA3, pS14-8(LEU2)

Y4129 MATa , MET3pr-HA-CDC20::URA3, pS14-8(LEU2)

Y4201 MATα, tor1-1, fpr1::NAT, RPL13A-2xFKB12::TRP1, scc1-

FRB::HIS3, pS14-8(LEU2)

Y4059 MATα, tor1-1, fpr1::NAT, RPL13A-2xFKB12::TRP1, brn1-


57


Y4199 MATa , pRS316(URA3), (S288c background)

Y4235 MATα, tor1-1, fpr1::NAT, RPL13A-2xFKB12::TRP1, scc1-

FRB::HIS3, smc2-FRB::kanR, pS14-8(LEU2)

Y4236 MATa , smc2-8, pRS316(URA3)


FRB::HIS3, pRS316(URA3)

Y4259 MATa , leu2Δ::kanR, RCIII-SUP11-LEU2-3ARS

Y4264 MATa , leu2Δ::kanR, brn1-9::TRP1, RCIII-SUP11-LEU2-3ARS

Y4327 MATα, tor1-1, fpr1::NAT, RPL13A-2xFKB12::TRP1,

leu2Δ::kanR, brn1-FRB::HIS3, RCIII-SUP11-LEU2-3ARS

Y4332 MATα, tor1-1, fpr1::NAT, RPL13A-2xFKB12::TRP1, pS14-

8(LEU2)


FRB::HIS3, pRS316(URA3)

Y4334 MATa , tor1-1, fpr1::NAT, RPL13A-2xFKB12::TRP1, top2-


2.1.2 Yeast strain creation, mating and tetrad dissection

Strains were constructed by transformation with the appropriate DNA

integration fragment designed for gene knockout or tagging. Affinity epitope tags

were fused at the gene endogenous loci for Western blot detection, or a 3xGFP

and mRFP cassette for detection by fluorescent microscopy, using polymerase

chain reaction products (Bähler et al., 1998, Knop et al., 1999). In order to cross to

strains, mating was induced through incubation of opposite mating type yeast

strains on YPD plates at 25°C for at least 6 hours. Diploids were then selected by

re-plating the crossed strains onto appropriate selective media and grown again on

YPD overnight. Diploid cells were sporulated on sporulation media (100mM

CH3COONa, 20 nM NaCl, 25 mM KCl, 1.5 mM MgSO4 and 1.5% w/v agar) until

tetrads appear. To break their asci, spores were treated with Zymolase T-20 (MP

Biomedicals) for 10 minutes at 30C. From each ascus, four spores are released

which were then dissected using a Singer-MSM micromanipulator. Finally, spores


58

are incubated at 25°C until colonies formed and from these colonies can then be

re-streaked onto the appropriate selective media to determine the genotypes of the

progeny generated.

2.1.3 Yeast media, cultures and synchronizations

Cells were grown in YP supplemented with 2% w/v glucose (YPD) or 2%

w/v raffinose/galactose (YP-Raff/Gal). Yeast cells of a mating type were arrested in

G1 with the mating pheromone α-factor. To arrest cells in G1, early log phase

cultures (OD600 = 0.1) was treated with 0.5 µg/ml α-factor (provided by peptide

services, Cancer Research UK). Cells of α mating type were synchronised using

0.04 µg/ml a-factor (provided by peptide services, Cancer Research UK), as

described (O'Reilly et al., 2012). One hour after the initial addition of α-factor or a-

factor, the same amount is added to the culture. Within two hours complete

synchronization of the culture in G1 is achieved. Cell cycle arrest was determined

both cytologically by the appearance of a characteristic ‘shmoo’ and by FACS

analysis of DNA content. G1 arrested cells were collected on a membrane filter

(Schleicher & Schuell, ME28, 1.2mm) using a filtration apparatus (Millipore). Cells

were washed with at least five times the initial culture volume of YPD before being

release into fresh YPD media.

For arresting cells at metaphase, 5 µg/ml nocodazole was added to the

culture from a 2 mg/ml stock solution in DMSO. To metaphase arrest strains using

the repression of MET-Cdc20, cells were grown in YNB supplemented with 2%

glucose. To induce arrest, 2 mM methionine was added to the culture.

2.1.4 Anchor away strains and nuclear deplet ion

Anchor-away strains were created by FRB-tagging of the target gene in the

anchor away strain background as described (Haruki et al., 2008b). To deplete

nuclei of cohesin or condensin, rapamycin was added to the respective anchor

away strains at the time of their release from synchronization in G1.


59

2.1.5 Yeast transformations

Transformation of cells with plasmids, minichromosomes or PCR products

was performed using standard lithium acetate transformation. In brief, 50 ml of mid-

log phase culture was pelleted at 3,000 rpm for 5 minutes. The cell pellet is washed

with 1 ml distilled water, then washed with 1 ml 1X TEL (10 mM Tris/HC pH 7.5, 0.1

mM EDTA, 100mM Lithium Acetate) before being re-suspended in a final volume of

100 µl 1X TEL. 1 mg of either linearised DNA vector or PCR product was mixed

with 2 ml of a 10 mg/ml single stranded carrier DNA from salmon sperm and 600 µl

TELP (TEL plus 40% PEG 3350 or 4000). 50 µl of the cell suspension in TEL was

then added to this mix and vortexed for 10 seconds. The mixture is incubated at

25°C for 4 hours and afterwards heat shocked at 42°C for 15 minutes. The cells are

pelleted at 6,000 rpm for 2 minutes, washed with and re-suspended in 1M sorbitol

before being plated onto YNB agar plates lacking the auxotrophic amino acid used

for selection. Transformants were checked for correct integration via either western

blot analysis, Southern blot analysis or in the case of anchor away strains, by death

on rapamycin containing media.

2.2 General molecular biology techniques

All standard molecular biology techniques such as PCR, restriction enzyme

endonuclease digestion or bacterial plasmid purification, were carried out as

described in (Sambrook and Gething, 1989) or as per the supplied manufacturers

protocol.

2.3 Protein analysis techniques

2.3.1 Protein extract preparation

Cell extracts were prepared using the TCA method. 5-10 ml of mid-log

phase cell culture is collected and cells pelleted by centrifugation, 3,000 rpm, 5

minutes at 4°C. Cells are resuspended in 1 ml of 20% trichloroacetic acid (TCA)

and kept on ice until the end of the time course. Cells are spun down for 1 minute

at 13,000 rpm, then washed with 1 ml of 1M Tris-Base before being resuspended in


60

100 µl 2X SDS-PAGE loading buffer containing DTT. Samples are boiled for 2

minutes at 95°C, then 100 µl of 0.5mm glass beads (BioSpec Products, Inc) were

added and the cells broken using a FastPrep FP120 cell breaker (Bio101). To

separate the cell lysate from the glass beads, a few small holes are made in the

bottom of the cell breaker tube using a 27G needle and the tubes are placed inside

15 ml Falcon tubes before being spun at 3,000 rpm, 5 minutes at 4°C. The

collected lysate is then boiled at 95°C for 5 minutes and cleared by centrifugation at

13,000 rpm for 5 minutes before being loaded onto an acrylamide gel.

2.3.2 Chromatin pel lets

Collect 50 ml of mid-log phase culture and centrifuge at 3,000 rpm for 5

minutes at 4°C to pellet cells. If performing timecourse, resuspend cells in 50 mM

HEPES/KOH pH 7.5, 100 mM KCl, 1.5 mM MgCl2, 1 M sorbitol and keep on ice.

When time course is over and samples can be processed, resuspend cells in 3 ml

of 100 mM PIPES/KOH pH9.4, 10 mM DTT, 0.1% Na-Azide) and incubate for 10

minutes at room temperature. Then spin the cells for 2 minutes at 2,000 rpm and

aspirate supernatant. Cells are resuspended in 2 ml of 50 mM KPi pH 7.4, 0.6 M

sorbitol, 10 mM DTT and to each sample 4 µl of 20 mg/ml Zymolyase 100T (MP

Biomedicals) is added for spheroplasting of the cells. After spheroplasting, all work

should be done at 4°C. Cells are spun down for a minute at 4,000 rpm and then

washed with 1 ml of 50 mM HEPES/KOH pH 7.5, 100mM KCl, 2.5 mM MgCl2, 0.4

M sorbitol and then spun again for a minute at 4,000 rpm. Cells are resuspended in

an equal volume of buffer EB (50 mM HEPES/KOH pH 7.5, 100 mM KCl, 2.5 mM

MgCl2, 1 mM DTT, 20 µg/ml leupeptin, 2 mM benzamidine, 2 µg/ml aprotinin, 0.2

mg/ml bacitracin, 2 µg/ml pepstatin A and 1 mM PMSF). To this 10% Triton X-100

is added to a final concentration of 0.25% and then incubated for 3 minutes on ice

with occasional vortexing, which produces the whole cell extract. Prepare 100 µl

EBX-S (EB + 0.25% Triton X-100 + 30% sucrose) in separate Eppendorf tubes and

lay 100 µl of the whole cell extract onto the EBX-S. Spin for 10 minutes at 12,000

rpm, which will produce a white chromatin pellet, a clear sucrose layer and above

this, a yellow supernatant fraction.


61

2.3.3 SDS-PAGE electrophoresis and western blott ing

Protein samples were resolved on acrylamide/bis-acrylamide (37.5:5:1,

amresco) 375 mM Tris-HCl pH 8.8 and 0.1% SDS. Small proteins of less than 30

kDa were typically resolved on 10-12% gels and larger proteins over 100 kDa on

8% gels. A stacking gel was used on top of the separating gel and consisted of 125

mM Tris-HCL pH 6.8, 5% acrylamide/bis-acrylamide and 0.1% SDS.

For electrophoresis a current of 50 mA was applied in an electrophoresis

tank (CBS Scientific) using SDS-PAGE running buffer (25 mM Tris, 250 mM glycine

and 0.1% SDS). To follow electrophoresis progression, and to allow the later size

comparisons of proteins on the gel, a broad range pre-stained protein marker (New

England Biolabs) was used.

After completion of electrophoresis, proteins were transferred onto pre-

equilibrated nitrocellulose membranes (GE Lifesciences) using a wet-transfer tank

(Biorad) and transfer buffer (3.03 g/l Tris base, 14.1 g/l glycine, 0.05% SDS and

20% w/v methanol). To check that the transfer was effective, the membrane is

stained with Ponceau S solution (Sigma). Then the membrane is blocked for an

hour with a 5% milk solution (Marvel) in PBST (170 mM NaCl, 3 mM KCl, 10 mM

Na2HPO4, 2 mM KH2PO4, 0.01% Tween 20) at room temperature. Following this,

membranes were incubated with primary antibodies diluted in milk solution for one

hour at room temperature, or overnight in the cold room. Primary antibodies used

and their final concentrations were a-Pk clone SV5-Pk1 (1:5000, Serotec), a-HA

clone 3F10 (1:5000, Roche), a-HA clone 16B12 (1:5000, Covance) and a-tubulin

clone YOL1/34 (1:1000, Serotec). After primary antibody incubation, membranes

were then washed in an excess of PBST for 30 minutes. Horseradish peroxidase

(HRP) coupled secondary antibodies (anti-mouse or anti-rabbit, 1:5000,

Amersham) were then incubated with the membrane in PBST containing 5% milk

for a further hour. Again membranes were then washed three times with an excess

of PBST for 30 minutes before developing with ECL (Amersham) according the

manufacturer’s instructions.


62

2.4 Mini-chromosome purif ication, electrophoresis and

catenane quantif ication

1 gram of cells were collected at each time point, resuspended in ice cold 1

M sorbitol, 0.1 M EDTA pH 7.5, 0.02% sodium azide, and kept on ice until the end

of the timecourse. Cell pellets were then resuspended in 0.5 ml of the same buffer,

20 µl of 2.5 mg/ml Zymolyase 100T (MP Biomedicals) was added and the

suspension incubated for 1 hour at 37°C with mild agitation. Cells were collected by

centrifugation for 10 minutes at 13,000 rpm, supernatants removed by aspiration

and the pellets resuspended in 0.5 ml of 50 mM Tris/HCl pH 7.4, 20 mM EDTA.

SDS was added to a final concentration of 1% and samples were incubated at

65°C for 15 minutes. Then, 0.2 ml of 5 M potassium acetate was added and the

samples placed on ice for 1 hour. The precipitate was removed by centrifugation for

5 minutes at 14,800 rpm and the supernatants collected. The centrifugation step

was repeated until the supernatant fractions were clear of any debris. 2 volumes of

100% ethanol were now added, and after 5 minutes the DNA was collected by

centrifugation for 1 minute at 13,000 rpm. The supernatants were aspirated and the

pellets allowed to air dry. Once dry, pellets were carefully resuspended in 0.3 ml of

TE, pH 7.4 and 50 mg/ml RNase A added and incubated for 30 minutes at 37°C,

before addition of 100 mg/ml proteinase K for further 30 minutes. Now, 12.6 µl of 5

M sodium chloride was added, and the DNA precipitated by addition of 0.63 ml

100% ethanol. Samples were again centrifuged, supernatants aspirated and the

pellets air-dried. After resuspension in TE, the DNA concentration was measured

(NanoDrop, Thermo Scientific) and adjusted prior to gel electrophoresis.

Samples were resolved on 0.5% agarose/TAE gels at 1.0 V/cm for 48 hours

(pRS316 and pS14-8) at room temperature, or at 0.8 V/cm for 24 hours, followed

by 2.2 V/cm for a further 24 hours (RCIII). Southern transfer was carried out using

capillary blotting onto a positively charged nylon membrane (Hybond-N+, GE

Healthcare) as per (Southern, 1975). The membrane was probed with random-

prime 32P-labeled probes against the AmpR gene (pS14-8 and pRS316) or against

LEU2 (RCIII). The blots were exposed to PhosphorImager screens (GE

Healthcare) that were scanned using a Storm 860 Molecular Imager. Band

intensities were analyzed and quantified using linear background subtraction and

automatic band detection in ImageQuant.


63

Enzymes used to verify the nature of the observed bands were XhoI and PmlI

(New England Biolabs), calf thymus topo I (Invitrogen), human recombinant topo I

and human topo IIa (both Topogen). When statistical analyses have been

performed, these have been done via paired t-tests using Prism software.

2.4.1 Catenation assay minichromosomes

The centromeric plasmid pRS316 (Sikorski and Hieter, 1989) and pS14-8

(containing a genomic region surrounding RAD5 in the centromeric plasmid YCp70

(Aguilera and Klein, 1990), a gift from A.-M. Farcas and K. Nasmyth) were

introduced into yeast using standard lithium acetate transformation. The ring

chromosome III (RCIII) contains 61 kb surrounding centromere III, including

ARS307, 308 and 309 as well as the wild type LEU2 gene (Dershowitz and Newlon,

1993). To identify RCIII by Southern blotting, the leu2-3.112 gene on the authentic

chromosome III in the host strain was replaced with a kanR marker (Wach et al.,

1994).

2.4.2 Zymolyation of yeast cel ls in agarose plugs and Pulse Field Gel

Electrophoresis (PFGE)

Collect approximately 108 cells and fix in 70% ethanol overnight at -20°C.

The following day, prepare the agarose plugs by mixing 1 ml of SEZ buffer (1M

sorbitol, 50 mM EDTA pH 8) with 0.02 g of LOM (low melting point) agarose and

heat to 65°C to melt the agarose. Cells are centrifuged for 1 minute at 13,000 rpm

and the 70% ethanol is aspirated off. Cells are then washed once in 1 ml of

resuspension buffer (1M Tris pH 7.5, 1.2M sorbitol, 0.5M EDTA) before cells are

resuspended in 50 µl SEMZ buffer (1M sorbitol, 50 mM EDTA pH 8, 28mM

mercaptoethanol, 3 mg/ml Zymolyase 100T (MP Biomedicals)). Cells in the SEMZ

buffer are then placed on a shaking heat block set at 37°C for 1 hour. After this

time, the cells are mixed with the melted LOM agarose (with 3 mg/ml Zymolyase

100T added) and pipetted into plug moulds for cooling and setting. Once plugs are

set, they are then incubated for 1 hour in a 37°C shaking heat block in EST buffer

(10 mM Tris pH 8, 100 mM EDTA pH 8, 1% sarcosyl). After an hour, the EST buffer


64

is removed and replaced with 50 mM Tris pH 7.8 and 10mg/ml RNAse A and left at

37°C overnight. The following day the buffer is removed and replaced with ESP

buffer (1% SDS, 1 mg/ml proteinase K, 0.5M EDTA pH 8) and incubated at 50°C

for 2 hours. Finally the buffer is changed to 1X TE and plugs can be stored at 4°C.

Prior to loading, plugs should be washed 3 times for 20 minutes each with 0.5X

TBE or 1X TAE buffer depending on the electrophoresis protocol being used. For

the last 20 minutes prior to loading, the plugs should be kept on ice.

Plugs are then loaded into 0.5% agarose gels and resolved with via PFGE.

In our protocol, we used 1X TAE as the running buffer. PFGE was performed on a

BioRad CHEF DR-III system at 14°C using the following program: Step 1: 24 hours

at 2 V/cm, 96° angle, 1200 seconds switch time; Step 2: 24 hours at 2 V/cm, 100°

angle, 1500 seconds switch time; Step 3: 24 hours at 2 V/cm, 106° angle, 1800

switch time.

2.5 Cell biology and microscopy

2.5.1 Cell cycle analysis using f low cytometry

To determine cell cycle progression by DNA content, 1 ml of culture was

pelleted and fixed in 70% ethanol for at least 2 hours at 4°C. Then cells are

resuspended in 50 mM NaCitrate containing 0.1 mg/ml RNAse A and put on a

shaking heat block at 37°C for an hour. After the hour has passed, proteinase K is

added to a final concentration of 0.1 mg/ml and the mixture left on the shaking heat

block for a further hour. Cells are then pelleted and then resuspended in 50 mM

NaCitrate containing 50 µg/ml propidium iodide. Cells are sonicated (Sanyo,

Soniprep 150) to eliminate clumping and then are analysed on a FACScan (Becton

Dickinson), with final data and graph generation prepared with CellQuest software.

2.5.2 Bi-nucleate cel l counting

The fraction of binucleate cells, an indication for progression through mitosis,

was scored using propidium iodide-stained cells. Scoring was repeated three times

over and an average taken.


65

2.5.3 In situ immunofluorescence

2 ml culture aliquots are collected with an OD600 of at least 0.2. The cells

are spun down and resuspended in 1 ml of ice cold formaldehyde buffer (100 mM

KPO4 pH 6.4, 0.5 mM MgCl2, 3.7% formaldehyde). Cells are fixed overnight in this

buffer at 4°C or can be process immediately by fixing for 2 hours at 30°C. Once

fixed, cells are washed once in 100 mM KPO4 pH 6.4, 0.5 mM MgCl2, then washed

once with 1 ml of spheroplasting buffer (100 mM KPO4 pH 6.4, 0.5 mM MgCl2,

1.2M sorbitol). After washing, cells are resuspended in 200 µl of spheroplasting

buffer plus 2 µl of β-mercaptoethanol and 2 µl of 20 mg/ml Zymolyase 100T (MP

Biomedicals) and incubated for 30 minutes at 30°C. Then cells are spun down for 2

minutes at 4,000 rpm and the supernatant is aspirated. Cells are washed with 0.5

ml of spheroplasting buffer before final resuspension in 0.2 ml of the same buffer. 5

µl of cells are then pipetted onto a polylysine coated 15 multi-well slide (MP

Biomedicals). Slides are washed for 3 minutes in methanol and fixed for 10

seconds in acetone before being blocked with blocking buffer (0.5% Bovine Serum

Albumin). Slides were then incubated with the relevant primary and secondary

antibodies in a humid chamber in the dark for an hour each. In between incubation

with each antibody, the slide wells were washed three times with blocking buffer

and four times before addition of a mounting anti-fade media with 0.1 µg/ml DAPI.

The antibody we used was α-tubulin clone YOL1/34 (Serotec).

2.6 Protein purif ications

2.6.1 Purif icat ion of S. cerevisiae topoisomerase II

Overexpression and purification of S. cerevisiae topoisomerase II followed a

published procedure (Worland and Wang, 1989). 6 hours after induction of topo II

expression by galactose addition, 0.5 grams of cells were collected, washed in

water, then resuspended in 200 µl lysis buffer (50 mM HEPES/KOH, pH 8.0, 150

mM KCl, 1 mM EDTA, 10% glycerol, 5 mM b-mercaptoethanol, 1 mM PMSF, and

an additional protease inhibitor cocktail (CompleteTM, Roche). Cells were broken

with acid washed glass beads to obtain greater than 90% cell lysis and the lysate

collected. A 1 ml phospho-cellulose column (Whatman P11, prepared according to


66

the manufacturer’s instructions) was equilibrated in lysis buffer. The cell lysate was

loaded in batch, then the resin was packed into a column and washed by gravity

flow using lysis buffer containing 200 mM and subsequently 400 mM KCl. Topo II

was eluted in lysis buffer containing 800 mM KCl. This fraction was diluted back to

200 mM KCl and bound to 300 ml Q-Sepharose beads (GE Healthcare). These

were then packed into a column and extensively washed with the loading buffer. A

step gradient with 1M KCl was used for elution. Topo II-containing fractions were

adjusted to 300 mM KCl and 10% glycerol for storage.

2.6.2 Purif icat ion of S. cerevisiae condensin

The 5-subunit condensin complex was overexpressed in S. cerevisiae as

previously described (St-Pierre et al., 2009). Cell extract containing 40 mg protein

in 6 ml of lysis buffer (20 mM HEPES/KOH pH 7.5, 150 mM KCl, 1 mM MgCl2, 5

mM β-mercaptoethanol, 10% glycerol, and protease inhibitors) was cleared by

ultracentrifugation. The supernatant was loaded onto 200 ml of an α-HA affinity

matrix (Roche). After binding, the resin was washed extensively in same buffer.

Condensin was eluted by incubation with 500 µl 1.6 mg/ml HA peptide in lysis

buffer over night at 4°C.

2.6.3 Interaction analysis between condensin and topoisomerase II

To carry out the interaction analysis of condensin with topo II, 25 units/ml

benzonase (Sigma) was added to the cell extract containing overexpressed

condensin. After ultracentrifugation, 400 µl aliquots of the extract were precleared

with 20 µl of IgG-coupled Dynabeads (Invitrogen) for 30 minutes at 4°C. The beads

were removed and 4 mg α-HA antibody (clone 16B12) was added to the

supernatant for 30 minutes at 4°C. Then, 20 µl of protein A-coupled Dynabeads

(Invitrogen) were added for a further 30 minutes. Finally, condensin-bound or

control beads were incubated in the presence of 400 ng purified S. cerevisiae topo

II for 30 minutes at 4°C. After washing, bound protein was elution in SDS-PAGE

loading buffer.


67

2.7 In vitro assay techniques

2.7.1 kDNA decatenation assays

To assess their decatenation activity, 10 ng or 100 ng of purified yeast topo

II, or 0.1 units of E. coli topo IV and human topo II (Topogen), were incubated with

100 ng of kinetoplast DNA (Topogen) in a 20 ml reaction containing 50 mM

Tris/HCl pH 7.8, 150 mM potassium acetate, 6 mM magnesium acetate, 5 mM b-

mercaptoethanol, 0.1 mg/ml BSA and 2 mM ATP. To assay the effect of condensin,

reactions were incubated with the indicated amounts of condensin for 30 minutes at

37°C in the absence of topoisomerase. Then topoisomerase was added and the

reaction incubated at 37°C for further 10 minutes. Reactions were stopped by

addition of an equal volume of stop buffer (2% SDS, 80 mM EDTA, 600 mM NaCl)

and the samples resolved on a 1% agarose/TAE gel. Gels were stained with

ethidium bromide and band intensities were quantified using ImageQuant software.

Chapter 3 Results

68

Chapter 3. Results: Catenane Behaviour and

Resolution

3.1 Development of the catenation assay

As discussed in the introduction, there are two aspects to chromosome

resolution, the cleavage of proteinaceous links and the resolution of topological

links between sister chromatids. It is only once both of these are complete that the

mitotic spindle can successfully pull apart the sister chromatids to the opposite

poles. Therefore in order to study the formation and resolution of sister chromatid

catenanes during the cell cycle, we employed circular yeast minichromosomes of

various sizes in a catenation assay that we tailored to our needs. The assay was

based on a similar protocol originally used to study whether the cohesin ring

physically entraps DNA, holding it together prior to seperase-cleavage and

anaphase onset (Ivanov and Nasmyth, 2007). The assay is a powerful tool for

tracking topological links through the cell cycle, because the minichromosomes,

which unlike linear DNA, maintain any topological links once purified. Moreover, it

has long been observed that larger genomes behave almost exactly like circular

DNA because of the existence of domains whose fixed ends act by preventing any

free rotation (Worcel and Burgi, 1972). However, in order to most closely model the

endogenous chromosomes, we chose to use minichromosomes for our assay that

retained many features of natural yeast chromosomes, including centromeres,

replication origins and actively expressed yeast chromosome arm sequences. Thus,

the minichromosomes would replicate and segregate with the same timing as the

native chromosomes, mimicking their behaviour.

The first substrate we chose to use in our assay was pS14-8, a 21.2 kb

centromeric mini-chromosome that contained a section of chromosome XII arm

sequence around the RAD5 gene (Figure 3.1). We chose to first use such a large

substrate based on prior reports that smaller plasmids are too quickly resolved of

their topological linkages by topoisomerase II to be observed in a normal

timecourse (Baxter et al., 2011), thus maximising our chances of observing

catenanes. For full details on the final protocol that we used, please see chapter

2.4 of the materials and methods. However, unless otherwise detailed, our

Chapter 3 Results

69

timecourses all adhered to the same general following framework: our yeast strain

of interest was transformed with our assay minichromosome and cultures grown

overnight. After dilution to the same optical density (OD) the following morning,

cultures were arrested in G1 through addition of the relevant mating pheromone.

Once arrested, cells were washed and released into pheromone-free media that

would either be shifted to 37°C or contain rapamycin (depending on the method of

protein inactivation). Cells would progress through the cell cycle before re-arrest

back in G1. From the moment cells are released into pheromone-free media,

aliquots for DNA purification are taken every 20 minutes as well as aliquots for

FACS analysis. After the timecourse is complete, the DNA is purified from the cells

and resolved on an ethidium bromide free gel at low voltage. To visualise the

minichromosome we performed a Southern blot and probed against a

minichromosome-specific gene. The end result is a blot showing several bands,

each representing a different topological isoform of the minichromosome in

question.

Chapter 3 Results

70

Figure 3.1 Map of minichromosome pS14-8

Chapter 3 Results

71

3.1.1 Optimisation of the catenation assay

Optimising the assay was a difficult process owing to the size of the plasmid.

Size is an issue because as the plasmid size increases, its electrophoretic mobility

decreases and this applies to all of its topological forms. This means that when

running the DNA on the agarose gel, it can be difficult to cleanly segregate different

isoforms into distinct bands, as they can all run with a similar speed. Initial blots

were difficult to interpret owing to how close bands ran together, making it difficult

to differentiate between them. In Figure 3.2, the first three lanes contain the purified

DNA from a wild type strain containing pS14-8 (Y2665). In comparison to DNA

purified from a wild type strain without pS14-8, we could clearly see some bands

specific to the minichromosome, yet with poor clarity. By careful troubleshooting of

the running conditions, such as the percentage of agarose used and the voltage

applied to the gel, a set of conditions was reached that allowed for improved

segregation of the minichromosome’s topological isoforms. We found that by using

a very low percentage agarose gel (0.5%) and running the samples very slowly

over a long period of time (1.0V/cm for 48hrs), blots were produced that were

generated greater spread of isoforms allowing for easier identification (Figure 3.3).

However looking at the blot, while the bands do segregate more clearly,

there persisted another problem. Despite the specificity of the probe for the AmpR

gene, it became apparent that the first probe we had produced was also weakly

labelling the genomic DNA. This can be seen as a large smeary band in the control

lane where DNA was prepared from wild type cells (K699) that contained no

plasmid or AmpR gene. Unspecific labelling of the genomic DNA was further

confirmed by staining gels after running with ethidium bromide to observe the

genomic band. Indeed, the distance travelled by this band and the distance

travelled by the smeary band on our Southern strongly suggested improper

labelling of the genomic DNA by our probe. Therefore in order to remove this

background labelling, we began testing a set of three new probes (labelled A, B

and C) that we designed against the AmpR gene that varied in length (Figure 3.4).

Chapter 3 Results

72

Figure 3.2 Early electrophoresis condit ions showed poor isoform

resolut ion

DNA purified from wild type cells containing pS14-8 (Y2665) is resolved electrophoretically and detected via Southern blot. Initially, running conditions were 5 V/cm for 3 hours on a 0.75% agarose gel.

Figure 3.3 Background labell ing of genomic DNA

By running samples for longer and at lower voltages (1.0 V/cm for 48 hours on a 0.5% agarose gel), a better spread of isoforms is obtained. However, our initial probe was not fully specific against the AmpR gene and labelled the

Chapter 3 Results

73

genomic DNA as seen in the lane where only DNA purified from wild type (K699) cell is run.

Figure 3.4 Specif ic i ty comparison of dif ferent AmpR probes

Three new probes specific for the AmpR gene were tested: probe A (length of probe = 469bp), probe B (524bp) and probe C (297bp). Probes were tested on DNA purified from wild type cells either containing pS14-8 (Y2665) or not containing pS14-8 (K699). Additionally, they were tested against the purified pS14-8 plasmid alone.

Chapter 3 Results

74

We used these three new probes to label either purified DNA samples from

Y2665, K699 or purified pS14-8 (i.e. no genomic DNA present) to compare the

intensity of their labelling as well as whether they had any non-specific labelling of

the genomic. Examination of the blots showed that the new set of probes all

seemed to be entirely specific for the AmpR gene with little to no affinity for the

genomic DNA, especially as when compared with our original probe as used in

Figure 3.2 and 3.3. However, they did not all label the AmpR gene with the same

intensity, and we determined that probe B labelled the most clearly and strongly. As

such, for all future experiments where we wished to target AmpR, we used probe B.

3.1.2 Identi fy ing the minichromosome’s dif ferent topological isoforms

Having optimised the conditions for resolving the DNA on the agarose gel so

that we could easily observe all topological isoforms, as well as having produced a

highly specific probe; we could start to use the assay to study the behaviour of

catenanes during mitosis. Having transformed pS14-8 into a wild type strain

(Y2665), we synchronised the culture in G1 using the relevant mating pheromone,

before washing the culture and releasing the cells from their arrest. Cells were

allowed to progress through one cell cycle before being rearrested in G1.

Throughout this timecourse, aliquots of cells were taken every twenty minutes and

were processed to purify the DNA. This DNA was then resolved on an agarose gel

at low voltage and a Southern blot performed, which was then probed against

AmpR using probe B (Figure 3.5).

Examining the blot, we can see a characteristic pattern of bands that

represent the different topological isoforms of the minichromosome pS14-8. While

two abundant bands are consistent throughout the cell cycle, it is clear that three

new bands become apparent at the time of S-phase that then disappear again in

mitosis. In order to identify which isoform each band represented, we performed a

series of enzyme digests upon our purified DNA (Figure 3.6). The first enzyme

used was Xho1, a restriction enzyme that only cuts pS14-8 once, hence linearizing

it. Indeed from the gel, one can see that upon incubation with Xho1, all isoforms

collapsed into a single linear (L) band. Linear DNA is present in all our purified DNA

Chapter 3 Results

75

preps, most likely as an unavoidable artifact of DNA shearing during the DNA

purification process. The next digest performed was with topoisomerase I (topo I),

an enzyme that is capable of relaxing supercoiling in DNA through the transient

‘nicking’ of a single strand of DNA, allowing the DNA helix to rotate and release

tension (see chapter 1.5.1). By treatment with this enzyme, bands representing

supercoiled isoforms of pS14-8 disappeared, with their intensity shifting to two

bands likely representing their relaxed forms, relaxed monomers (RM) and relaxed

catenanes (RC). The final enzyme treatment was with topoisomerase II (topo II),

which differs from topo I in that it can create transient double-strand breaks,

allowing the passage of one DNA strand through another, before re-ligating the

DNA (see chapter 1.5.2). This means that topo II cannot only relax supercoiling

(like topo I), but also can resolve topological links between different DNAs.

Consequently, we saw that treatment with topo II resulted in any bands

representing catenated pS14-8 disappear, leaving one intense band representing

the relaxed monomer form of pS14-8. However, a smear is seen underneath this

band and this likely represents partially supercoiled monomers (PSM). This is a

result of topo II’s bi-directionality, meaning that it can both introduce and remove

supercoiling and catenation. In the lane where the DNA is treated with topo II, the

smear represents monomers of pS14-8 where an equilibrium distribution of partially

supercoiled monomers has been established. The reason that all the monomers

are not partially supercoiled and we have a strong relaxed monomer band, is

because the relaxed monomer band contains plasmids that have likely been

‘nicked’, i.e. had a single strand of DNA broken, either as a result of shearing

during pipetting or as consequence of the enzyme treatment. If nicked, plasmids

are unable to sustain any supercoiling because any tension introduced is

immediately lost. Topo II can also introduce topological links, but the reason we

see no catenanes is because the equilibrium is shifted very much towards

decatenation. For topological links to be introduced between different plasmids,

they need to be physically very close together for the reaction to occur. At their low

concentration and with no proteins holding them together, it is therefore very

unlikely for links to be introduced. For those plasmids that are catenated, once

decatenated by topoisomerase II, the now individualised plasmids will separate

spatially from each other rapidly owing to Brownian motion.

Chapter 3 Results

76

Figure 3.5 Wild type cel ls display a dist inct isoform patterns

Wild type cells containing pS14-8 (Y2665) were synchronised in G1 and then released, allowed to progress through one cell cycle before re-arrest. The appearance of new topological isoforms occurs when DNA replication begins, which then disappear in mitosis. FACS analysis and counting of binucleate cells confirms the synchrony of the cells and the specific timing of the appearance and resolution of these transient isoforms.

Figure 3.6 Enzyme digests al low for isoform identi f icat ion

Enzyme treatment revealed the identity of isoform represented by each band; relaxed catenanes (RC), mixed catenanes (MC), relaxed monomers (RM), partially supercoiled monomers (PSM), supercoiled catenanes (SC), linear (L) and supercoiled monomers (SM). Catenated isoforms are marked with a star.

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Having performed these enzyme treatments, we could confidently identify

which topological isoform each band on the Southern blot represents. There are

three bands detectable that represent catenated species and what is immediately

striking when examining the wild type timecourse is how these catenated forms

appear and then disappear with a specific pattern over the course of the cell cycle.

Comparing the blot with the FACS profile, one can see that catenanes appear in S-

phase when DNA replication occurs, matching the prediction made by (Sundin and

Varshavsky, 1980) 30 years ago that catenanes are replication dependent.

Furthermore, one can see that these catenanes are then resolved later in the cell

cycle, approximately around the time of anaphase onset. We will investigate further

the exact timing of this resolution in later experiments.

3.1.3 Catenation is quanti f iable at each t ime point

To be able to observe catenanes using pS14-8, and to measure the specific

timing of their appearance and resolution, was very exciting. However before

investigating the behaviour of these catenanes any further, we required a way to

quantify the relative amount of catenanes present at each time point taken during

the time course. This would allow us to then quantify the effect certain mutants or

nuclear depletions had on catenation creation, resolution and behaviour. We

accomplished this through the use of ImageQuant imaging software, the final

protocol for which is described in the materials and methods (chapter 2.4).

There were many ways in which to calculate the intensities of different

peaks, including factors such as band detection and boundary determination. As far

as possible, all band detections and subsequent calculations was performed by the

program to minimise external influence on the data. However, correctly determining

how to apply the subtraction of background was an area we had to optimise. Figure

3.7 shows a comparison of two methods of background deduction: the rolling ball

and linear background subtraction methods. The rolling ball method is a computer

determined background subtraction that ‘rolls’ an imaginary ball of set diameter

along the underside of the lane profile to determine the background. However, one

can see that this technique is not suited to the types of profiles generated by our

data, with steep gradients seen at the start and end of the gel runs. This results in

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abnormally large quantification values for the first and especially the last band. An

improvement on this is the linear background subtraction along the visible

background seen in the lane profile. This deducts the background evenly and fairly

from each band’s quantification. Indeed in Figure 3.7 we can see the differences

between the two methods, which despite both quantifying the same wild-type

timecourse, it is clear that the linear background subtraction method generates

quantification data that best represents what can be seen on the Southern blot.

Once we had optimised the quantification protocol, we were able to start

putting numbers to the observations noted in the catenation assays. Returning to

our wild type timecourse (Figure 3.5), we quantified the amount of catenanes

present to produce a graph showing the amount of catenanes present as a

percentage of the total minichromosome in each lane (Figure 3.8). Therefore

quantification reveals that during DNA replication, the percentage of catenanes

peaks at around 55% before being resolved in mitosis.

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Figure 3.7 Comparison of background deduction techniques

The ImageQuant software can subtract the background via different algorithms such as (A), the ‘rolling ball’ method (radius = 200) or (B), a linear best fit. The above profile graphs represent individual lanes, with each number peak corresponding to a different isoform (1 = RC, 2 = MC and so on). The pink line represents the background threshold, such that the integrated values from under that line are subtracted from the final quantification values.

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Figure 3.8 Catenane quanti f icat ion of wi ld type cel l cycle

Quantification of the Southern blot seen in Figure 3.5, where synchronous wild type cells (Y2665) were allowed to undergo one cell division. A peak of approximately 55% total catenanes was reached during DNA replication.

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3.2 Spontaneous and assisted minichromosome

decatenation:

With the assay established, we were keen to start examining the effects on

catenation by depleting particular components of the mitotic assembly. For most of

our experiments involving the inactivation of a target protein, this has been

achieved through use of both temperature sensitive (ts) alleles and nuclear

depletion via the anchor away technique (see chapter 1.7). For the relevant

controls for both techniques, please see the next chapter 3.2.2.

The pS14-8 mini-chromosome harbours the efficient ARS1 early replication

origin and is expected to replicate and equally segregate during every cell cycle. If

catenation does indeed arise as a consequence of DNA replication, we would

expect to at least transiently observe all of the mini-chromosome to be present in

catenated forms. However, as seen in our wild type time course (Figure 3.5), the

peak was never higher than 60%. This could be because replication does not

always lead to catenation between replication products. Alternatively, it could be

that all replication products are catenated, but that some sister chromatids are

resolved quickly after replication, even before cells enter mitosis. To distinguish

between these two possibilities, we repeated the time course analysis but

inactivated the major endogenous decatenating activity of topo II using both the

temperature sensitive top2-4 mutation (Y4007) (Holm et al., 1985) and nuclear

depletion via anchor away (Y4334) (Figure 3.9). With both types of depletion, we

observed that upon entry into S-phase, virtually all of the minichromosomes

accumulated as catenated species following DNA replication. The fact that almost

no individual plasmids reappear supports the fact that without topoisomerase II,

decatenation does not occur. Additionally, it was reported that without

topoisomerase II, DNA replication does not progress to completion (Baxter and

Diffley, 2008) an idea supported by the observation of plasmid fragmentation during

anaphase as represented by the smear underneath the linear band. When

comparing the ts mutant phenotype with the anchor away, the former appears to

have a stronger phenotype. We believe that given the vast abundance of

topoisomerase II present in the nucleus, anchor away may not be fully able to

completely deplete the nucleus of the enzyme, or do so in a timely enough manner,

while all the ts mutant proteins will be inactivated upon temperature shift.

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Figure 3.9 Topoisomerase II inactivat ion results in maximum catenane

formation

Topoisomerase II is inactivated through either ts mutation (Y4007) or nuclear depletion via anchor away (Y4334). Both strain cultures are arrested in G1 at

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25°C before being washed and released into either media at 37°C or into media containing rapamycin.

Therefore, inactivation of topoisomerase II results in almost complete

conversion of the plasmid monomers during replication to catenated forms,

suggesting that S-phase results in all pS14-8 minichromosomes undergoing

replication and becoming at least transiently catenated.

To estimate which portion of sister minichromosomes are resolved before

progression through mitosis, we quantitatively assessed the catenation status of

pS14-8 in wild type cells (Y2665) arrested in mitosis by nocodazole treatment

(Figure 3.10). After a peak of catenanes during S-phase, over half of sister

minichromosomes appeared to be resolved in the mitotic arrest, with approximately

40% of the minichromosomes persisting as catenanes. Thus substantial

minichromosome decatenation by topo II takes place independently of progression

through mitosis. We then compared a mitotic arrest by depletion of the Cdc20 co-

activator of the anaphase promoting complex (Y4129) (Figure 3.11). In this strain

the endogenous Cdc20 promoter has been replaced such that the addition of 2 mM

methionine will stop Cdc20 expression, resulting in a metaphase arrest. In contrast

to nocodazole treatment, a mitotic spindle is formed in these cells that exerts

tension aimed at separating sister minichromosomes (Tanaka et al., 1999). As a

result, after similar catenane levels during S-phase, only about 20% catenated

molecules persisted in the arrest. This suggests that sister chromatid movement

away from each other, due to tension from the mitotic spindle, promotes their

decatenation.

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Figure 3.10 Nocodazole arrests results in catenane persistence

Wild type cells (Y2665) are released from G1 arrest into media containing nocodazole, causing cells to arrest in metaphase. After a replication peak of catenanes, the arrest results in the majority remaining unresolved.

Figure 3.11 Spindle presence during arrest promotes decatenation

Like in Figure 3.10 synchronised cells are arrested in metaphase, but here via depletion of Cdc20 using strain Y4129. This results in the mitotic spindle being present, which results in fewer catenanes persisting.

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Up until mitosis, movement of sister chromatids away from each other is

restricted by the cohesin complex that maintains sister chromatid cohesion. We

therefore addressed whether removal of cohesin changes catenane maintenance

following DNA replication. After either inactivation via ts allele or nuclear depletion

via anchor away, the abundance of catenanes during the cell cycle was markedly

reduced (Figure 3.12). Even during S-phase, less than 20% of the pS14-8

minichromosomes were detected as catenated species. This suggests that sister

chromatid proximity, afforded by the cohesin complex, protects catenation. In the

absence of cohesin, sister chromatids move apart and are readily decatenated by

topo II. The conclusion that cohesin protects sister chromatid catenation was

recently reached independently (Farcas et al., 2011). Quantification of the

catenanes for all the above time courses can be seen and compared in Figure 3.13.

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Figure 3.12 Inactivation of cohesin results in fewer observable catenanes

Cohesin is inactivated through either ts mutation (Y4029) or nuclear depletion via anchor away (Y4201). Both strain cultures are arrested in G1 at 25°C before being washed and released into either media at 37°C or into media containing rapamycin.

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Figure 3.13 Catenane quanti f icat ion for dif ferent mitot ic inactivations

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3.2.1 Temperature’s effect on catenation and isoform distr ibution

In the previous chapter, we have been using two different approaches to

inactivate target proteins: protein inactivation via ts alleles and nuclear depletion via

anchor away. Both of these procedures will inactivate the target protein but do so

by differing means, one requires a dramatic temperature shift, the other the

addition of the compound rapamycin. Given that both techniques alter the cellular

environment, we wanted to check that these depletion methods themselves did not

affect our catenation assay via the appropriate control experiments.

Initially, we examined the effect of temperature on our catenation assay. To

examine this, we released synchronous wild type cells (Y2665) from a G1 arrest

and allowed them to proceed through one cell cycle before re-arrest. This time

course was done at both 25°C and 37°C and the Southern blots were quantified

and compared (Figure 3.14). We can see that the pattern and distribution of bands

and their relative intensities is altered between the two different temperatures.

Moreover, a very slight amount of persistent catenation was detected in the 37°C

samples suggesting that the insult of the higher temperature could have some

slight effect on catenation, perhaps via impairment of topo II activity.

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Figure 3.14 Effect of temperature on catenane behaviour

Cells (Y2665) synchronised in G1, were released from their arrest into media either at 25°C or 37°C and allowed to progress through one cell cycle before re-arrest in G1.

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Following on from these observations, we also wanted to examine in more

detail the effect of temperature on the different topological forms. In the previous

results chapter 3.2, we found that a nocodazole arrest results in a catenation

equilibrium being established where all the different topological forms are present

and can clearly be seen. We repeated this experiment but this time performing the

experiment at 37°C instead of 25°C then proceeding to quantify and compare the

experiments (Figure 3.15). From this comparison, we made two interesting

observations; the first was that the overall amount of persistent catenation at either

temperature was very similar. The second was that the amount of supercoiled

species, both monomers and catenanes, was markedly reduced at higher

temperatures throughout the entire time course when compared to the levels seen

at 25°C. This is an unsurprising result, considering that increasing the temperature

will increase the kinetic energy of small molecules such as DNA. At higher

temperatures, collisions, interactions and reactions between molecules will occur

with greater energy. It is likely that this could translate to a higher occurrence of

DNA strand-nicking and as discussed earlier, plasmids that contain nicked DNA are

unable to sustain any supercoiling. In conclusion, these temperature comparisons

have shown that while increased temperature does affect the various ratios of

topoisomers present, the overall relative amount of catenation remains very similar.

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Figure 3.15 Effect of temperature on dif ferent isoforms

Wild type cells (Y2665) were synchronised in G1 and released into a nocodazole-induced metaphase arrest at either 25°C or 37°C. Quantification revealed that while overall catenation remained similar, supercoiled isoforms (both monomers and catenanes) were reduced at the higher temperature.

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3.2.2 The anchor away system does not effect catenation

When the new anchor away technique was published (Haruki et al., 2008a),

our lab quickly obtained the strain and modified it for our own ends by tagging our

proteins of interest with the FRB tag (see chapter 1.7 for an introduction to the

anchor away technique). Having tagged our target proteins, we wanted to check

that the parental strain itself, as well as the presence of rapamycin, did not exert

any influence on catenane behaviour. The anchor away strain has certain

mutations inherent to its design. Because rapamycin is toxic to wild-type S.

cerevisiae, the anchor away strains are made rapamycin-resistant through mutation

of TOR1 (tor1-1). Additionally FPR1 (Δfpr1) is deleted, as it is the yeast homolog of

the human FKBP12 gene that encodes the most abundant FK506 and rapamycin

binding protein in S. cerevisiae. This is necessary to reduce competition between

Fpr1p and the anchor-FKBP12 construct for binding to the FRB domain (Heitman

et al., 1991). Therefore given the modifications of the strain, plus the exposure to

the usually toxic rapamycin, we needed to ensure that neither had any effect on the

creation and resolution of catenanes. To test this, we performed a time course

using an anchor away ‘parental’ strain (Y4332) where no target protein has been

FRB-tagged. These cells were synchronised and released into media containing

rapamycin and allowed to progress through one cell cycle (Figure 3.16). As can be

seen, neither the strain nor the presence of rapamycin appears to have any

influence on catenation and the pattern of topological forms looks the same as with

our wild-type strain (Figure 3.5).

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Figure 3.16 Anchor away and rapamycin do not affect catenation

Strain Y4332, which does not contain any target FRB tagged protein, was synchronised in G1 and then released into media containing rapamycin. After progressing through one cell cycle, cells were re-arrested back in G1.

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3.3 Catenation of smaller minichromosomes is observable

in wild type strains

Having established the assay using the minichromosome pS14-8, and

having seen how clearly we were able to observe catenanes, we wanted to check

that our results were not unique to this one particular plasmid. Previous catenation

studies involving smaller minichromosomes, specifically pRS316 (4.88kb) (Figure

3.17), have reported that under wild type conditions, catenanes were resolved too

quickly during mitosis to be detected and could only be observed through

inactivation of endogenous topo II (Baxter et al., 2011). However, given the clarity

of our results with pS14-8, we decided to test a smaller minichromosome in the

assay to see if this was indeed the case. We also used pRS316 and apart from

changing the electrophoresis running conditions, used the same protocol for

purification and resolution of the DNA as with pS14-8. We performed a time course

using WT cells (Y4199), quantified the data and were able once again to observe

the appearance and then resolution of catenanes during the cell cycle (Figure 3.18).

To confirm the detection of catenanes, enzyme digests were performed on the

DNA samples to confirm what topological forms each band represented. Digests

with topo I and II revealed that the top two bands with the lowest electrophoretic

mobility indeed represented catenated forms. While the top band contains relaxed

catenanes, the second catenated band most likely contains a mix of both

supercoiled and mixed catenanes. While with pS14-8 these two forms ran

separately, it seems most likely that with the smaller pRS316, the supercoiled and

mixed catenanes have a similar electrophoretic mobility. The linear form of pRS316

was not detectable in our samples, suggesting that these smaller DNA molecules

are less susceptible to shearing during purification than their larger counterparts.

Comparing the results with pS14-8, it was reassuring to obtain similar results with

the much smaller minichromosome and additionally confirm that our catenation

assay was not particular to any one plasmid but rather representative of them all.

However, we can see from the quantification that the smaller pRS316 had a peak

catenation of under 30%, around half of the peak seen for pS14-8, suggesting that

most of the plasmid molecules were decatenated soon after synthesis. So while it

is very much possible to observe the catenanes of smaller plasmids under wild type

conditions, their smaller size seems to result in faster decatenation.

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Figure 3.17 Map of minichromosome pRS316

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Figure 3.18 Catenation detection and isoform identi f icat ion for pRS316

Catenanes can be detected in wild type cells containing pRS316 (Y4199). Cells are synchronised in G1 before release, and are allowed to progress through one cell cycle before re-arrest in G1. Enzyme treatments were used to determine which isoform each band represented; relaxed catenane (RC), supercoiled catenane (SC), mixed catenane (MC), relaxed monomer (RM), linear (L), partially supercoiled monomers (PSM) and supercoiled monomer (SM). Catenated isoforms are marked with a star.

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Chapter 4. Results: Condensin’s Inf luence on

Catenation

4.1 Condensin promotes completion of minichromosome

decatenation

Having established our catenation assay and studied some of the major

mitotic components, it was time to begin studying condensin directly. The first and

simplest approach to take was to inactivate condensin using one of the ts mutants

in our laboratory’s possession. Therefore, we performed timecourses with three

condensin ts alleles: smc4-1 (Y3954), brn1-9 (Y3939) and ycg1-10 (Y3940), each

containing the 21 kb minichromosome, pS14-8. In all cases, cells were

synchronised in G1 at the permissive temperature before being released into media

at the non-permissive temperature (Figure 4.1). Looking at the results for all three

mutants and comparing them with wild type cells shifted to 37°C, one can see a

subtle but constant phenotype of partial persistent catenation once mitosis is

complete. Quantification of the blots reveals that after peaking at approximately

50% catenanes 60 minutes after release from G1, the cells still retain 10-20%

catenanes total 180 minutes after release. When compared to wild type cells, that

have less than 5% catenanes upon re-arrest in G1, these results clearly show that

with condensin inactivated, there is a failure to completely resolve catenation. The

FACS profiles for all three condensin mutants also show a characteristic double

peak or ‘shoulder’ as the cells are re-arrested in G1 at the non-permissive

temperature. This is a characteristic condensin phenotype (D'Ambrosio et al.,

2008a) attributed to chromosome mis-segregation.

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Figure 4.1 Condensin ts mutants al l display a persistent catenane

phenotype

Condensin mutants ycg1-10 (Y3940), brn1-9 (Y3939) and smc4-1 (Y3940) are synchronised in G1 before release into media at the non-permissive temperature. For all ts mutants, persistent unresolved catenation is detectable when compared to wild type (Y2665) cells.

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Having obtained these results with the ts mutants, we were then very keen

to see if similar results were obtained via nuclear depletion of condensin using the

anchor away technique. Therefore, we analyzed catenane abundance in

synchronised cells following nuclear depletion of condensin using an anchor away

allele of its Brn1 subunit (brn1-aa (Y4059) Figure 4.2). Catenanes accumulated

during S-phase to comparable levels, irrespective of whether rapamycin was added

to deplete condensin. During G2 and mitosis catenanes started to be resolved

under both conditions but, in contrast to wild type cells that saw complete

decatenation, decatenation progressed more slowly in condensin-depleted cells

and approximately 15% catenanes persisted unresolved until the end of the

experiment. This was not because mitosis was delayed in the absence of

condensin. FACS analysis of the DNA content, as well as the budding index,

demonstrated that both cultures progressed through the cell cycle with similar

kinetics and all cells had returned to G1 by the end of the time course. Anaphase

bridges and chromosome missegregation were abundant after rapamycin addition,

as seen on the FACS profile and in anaphase cells stained with the DNA dye 4',6-

diamidino-2-phenylindole (DAPI; Figure 4.2), verifying that the depletion of

condensin was not just having some minichromosome-specific effect. Persistent

minichromosome catenation after condensin depletion was reproducibly observed

at similar levels in several biological repeats of this experiment (Figure 4.3).

Therefore, these results suggest that condensin promotes resolution of sister

chromatid catenanes and demonstrates that in the absence of condensin a fraction

of catenated minichromosomes persist until the time of cell separation by

cytokinesis.

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Figure 4.2 Condensin is required for complete decatenation

Condensin depletion causes part of the minichromosome catenanes to remain unresolved. Condensin was depleted via anchor away depletion of Brn1 (Y4059) before release from G1 arrest and allowed to proceed trough one cell cycle before re-arrest. The percentage of catenanes was quantified Immunofluorescence microscopy of cells stained with a α-tubulin antibody and the DNA dye 4',6-diamidino-2-phenylindole (DAPI) reveals anaphase bridges after condensin anchor away.

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Figure 4.3 brn1-aa tr ipl icates and quanti f icat ion

The experiment shown in Figure 4.2 was repeated 3 times and the percentage of catenanes at the indicated time points quantified. Only time points at 0, 60 and 180 minutes after release were taken. The mean and standard deviation are shown. Catenanes are marked with a star.

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If our circular minichromosome is a faithful model for the behavior of authentic

chromosomes, then the condensin requirement for complete decatenation could

explain anaphase bridges and chromosome missegregation in the absence of

condensin. One caveat to this conclusion is that if condensin was required to

resolve different, catenation-independent, linkages on authentic chromosomes,

then such linkages might indirectly hamper minichromosome decatenation. We

therefore addressed whether condensin contributes to chromosome decatenation

independently of promoting chromosome movement. To do this, we arrested cells

in mitosis using nocodazole, a situation where the mitotic spindle is disrupted and

therefore directed chromosome movement no longer contributes to sister chromatid

resolution. Cohesin was depleted from the nucleus using the scc1-aa allele, as

above, allowing spontaneous catenane resolution (Figure 4.4). Condensin

depletion in the same cells, via the brn1-aa allele, led to an increased population of

persisting catenanes. This suggests that condensin supports chromosome

decatenation independently of, or in addition to, promoting efficient chromosome

movement during anaphase. However, paired t-test analysis comparing the two

strains produces a P value of 0.0497, which only just implies a significant difference

at a confidence level of 95%. Following a similar line of thinking, we wanted to

confirm that condensin also supports decatenation independently of mitosis

progression. In Figure 4.5, smc2-aa MET3pr-cdc20 cells were arrested with

addition of methionine either in the presence of rapamycin or not. One can see that

when condensin is depleted, more catenanes persist, thus suggesting that the

mechanism by which condensin promotes decatenation is active prior to the

metaphase to anaphase transition. The effect of condensin here is more significant,

with a paired t-test analysis producing a P value of 0.0010, suggesting that there is

indeed a significant difference resulting from condensin depletion.

Taking these results together, we suggest that persistent catenanes in the

absence of condensin are a likely source of chromosome segregation failures.

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Figure 4.4 Condensin contr ibutes to decatenation independently of

chromosome movement

Cells of the indicated genotypes (Y4201, Y4235) were synchronised in G1 and released into media containing nocodazole and rapamycin. Catenanes were quantified and plotted with a comparison to arrested nocodazole wild type cells.

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Figure 4.5 Decatenation is active prior to anaphase

Cells of the indicated genotypes (Y4113) were synchronised in G1 and released into media containing methionine and either rapamycin or no rapamycin. Catenanes were quantified and plotted.

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4.2 The effect of minichromosome size

This clear observation of persistent catenation resulting from the inactivation

of condensin was stimulus to see if we observed a similar phenotype when

employing the smaller pRS316 plasmid. We began by investigating the effect of the

ts mutant smc2-8, in order to make a direct comparison to results from a previous

study (Baxter et al., 2011) where catenanes had not been seen upon shift to the

non-permissive temperature (Figure 4.6). In our timecourse we do clearly see

catenanes appear and it seems that a portion of these do persist. However, this

particular mutant had difficulty in release synchronously from the G1 arrest such

that we could not be certain that the catenanes observed 120 to 180 minutes after

release were not a result of late comers entering S-phase. The FACS profile is

particularly unclear for this strain (Y4236) making assessment of cell synchrony

difficult. Fortunately, we also had the anchor away method at our disposal (Figure

4.7). Nuclear depletion of condensin revealed a small delay in catenane resolution,

but the effect was less pronounced compared to the larger minichromosome.

However the FACS profile does show the distinct ‘shoulder’ upon re-arrest in G1

that is commonly seen in condensin mutants. By the time cells passed through

mitosis in the absence of condensin, less than 10% of catenanes persisted. This

suggests that condensin makes a contribution to plasmid decatenation, albeit small.

We note that if condensin-dependent positive supercoiling was required for

pRS316 decatenation, which has been observed in topo II depleted cells (Baxter et

al., 2011), we should have observed a greater resolution defect.

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Figure 4.6 smc2-8 releases poorly from G1 arrest

smc2-8 cells (Y4236) arrested in G1, washed and placed into fresh media at 37°C, release from the arrest asynchronously making the tracking of catenane appearance and resolution difficult.

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Figure 4.7 Condensin deplet ion has a smaller effect on pRS316

catenation

Cells of strain Y4333 were synchronised in G1 and released into media containing rapamycin before re-arrest. Catenanes were quantified and plotted.

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Compared to authentic chromosomes, pRS316 and pS14-8 are small in size

and both contain only one replication origin. Given that catenation results from the

meeting head-on of two replication forks during DNA replication, one can deduce

that the number of topological links between catenated pRS316 and pS14-8

plasmids are few. In addition once a link is removed, if the plasmids are not held

together by further links (or as we demonstrated, cohesin), then they will quickly

physically separate and are unlikely to be recatenated by topoisomerase II. Given

this, we hypothesised that condensin’s contribution to decatenation would be

greater in more complex topological landscapes than small minichromosomes. To

address this prediction, we adapted our catenation assay to use a much larger

substrate. In order to better represent the native chromosomes that this assay is

modelling, we studied a 61 kb ring chromosome (RCIII, Figure 4.8), consisting of a

61kb section of S. cerevisiae chromosome 3, including the centromere, which has

been excised then ligated to form a circular chromosome (Dershowitz and Newlon,

1993). In addition, this ring chromosome contained three origins of replication that

would result in an increased number of topological linkages being formed between

replicated DNAs during S-phase.

However, given the much larger size of this substrate, we found it much

more difficult to separate its topological isoforms using the same purification and

electrophoresis conditions as for pS14-8. Furthermore, owing to its much larger

size, the ring chromosome was more susceptible to shearing during DNA

purification. This resulted in a large fraction being linearised and creating greater

background signal that made band quantification difficult.

4.2.1 Visualiz ing a r ing chromosome’s (RCIII) dif ferent isoforms

In order to address the difficulties in resolving this much larger ring

chromosome, time was spent developing a new protocol to process the cell

samples. In this new protocol, cells were suspended in agarose blocks in which

they would then be lysed and processed to extract their DNA without any pipetting

involved at all, following a specific protocol as described in (Jain et al., 2010). It

was hoped that this gentler approach to DNA extraction would reduce or eliminate

linearization of RCIII. In addition, the DNA from the agarose plugs would then be

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resolved using Pulse Field Gel Electrophoresis (PFGE), a technique commonly

used for DNAs of a large size. However, despite extensive troubleshooting the best

results achieved (Figure 4.9) did not offer the resolution we hoped for. While it

appeared that we had been successful in minimising the amount of linear forms

generated (as none can be observed), the clarity of the different bands remained

poor and offered little advantage over our previous DNA purification protocol. As

such, we discontinued using PFGE and agarose gel plugs for DNA extraction and

instead redesigned our previous DNA purification protocol to minimise the number

of steps involving pipetting. In addition, we optimised the electrophoresis running

conditions to give us the best resolution of isoforms possible, as described in

chapter 2.4.

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Figure 4.8 Map of r ing chromosome RCIII

Figure 4.9 Resolut ion of RCIII by Pulse Field Gel Electrophoresis (PFGE)

Wild type cells (Y4259) were all metaphase arrested with nocodazole so that all isoforms would be present upon purification of DNA. Samples were digested in agarose plugs and DNA resolved by PFGE. However, despite eliminating shearing and linearization of RCIII, the resolution of individual isoforms is poor and are difficult to identify.

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4.3 Condensin’s decatenation role is more pronounced on

RCIII

Having tried PFGE and returned to our original (albeit optimised) purification

and electrophoresis protocol, we were able to find electrophoretic conditions that

revealed wild type catenane formation during DNA replication (Y4259) (Figure

4.10). We were able to observe a discrete band appearing below the relaxed

monomer (RM) band during DNA replication, that we identified as supercoiled

catenanes (SC) via enzyme treatment. Incubation with topo I relaxed these species

into a smear of relaxed and mixed catenanes (MC), probably of varied supercoiling

status, migrating above relaxed monomers. Their catenated nature was confirmed

by topo II treatment, which resolved these species into monomers. The general gel

background in the region of mixed and relaxed catenanes, however, prevented

their reliable analysis. In Figure 4.11 we see a quantification profile generated by

the ImageQuant software. Given that many of the catenated forms run as a smear

overlapping with bands representing monomers, the software is unable to correctly

quantify each topological form. Despite this setback, we proceeded to perform

several timecourse experiments using RCIII.

Focusing on the diagnostic supercoiled catenanes (SC), we observed their

appearance in S-phase, followed by resolution at the time of mitosis. As with the

smaller minichromosomes, only a fraction of the ring chromosome appeared

catenated following S-phase. This suggests that a sizeable portion even of this

large ring chromosome is decatenated rapidly following its replication. When we

blocked mitotic progression using nocodazole, a similar level of catenanes was

detectable during S-phase, but these were no longer resolved and persisted for an

extended period in the arrest (Figure 4.12). The same was observed after

condensin inactivation using the brn1-9 mutation (Figure 4.13). In this case cells

progressed through mitosis, yet catenanes following DNA replication persisted

without sign of resolution for the remainder of the time course. A similar result was

obtained following nuclear depletion of condensin (Figure 4.14). This suggests that

two pools of catenanes are formed in S-phase. One pool is readily resolved

following DNA synthesis, while a second pool of catenanes persists that is helped

by condensin for their decatenation. The condensin requirement for resolution of

this latter pool becomes more pronounced as chromosome size increases. In the

Chapter 4 Results

112

case of the 61 kb ring chromosome, resolution of persistent catenanes showed a

strict condensin requirement.

Chapter 4 Results

113

Figure 4.10 RCIII wi ld type t imecourse and isoform identi f icat ion

Wild type cells containing RCIII (Y4259) were synchronised in G1 and then released to progress through one cell cycle before re-arrest in G1. Enzyme treatments were used to determine which isoform each band represented; relaxed catenane (RC), supercoiled catenane (SC), mixed catenane (MC), relaxed monomer (RM), linear (L), and supercoiled monomer (SM).

Chapter 4 Results

114

Figure 4.11 RCIII Quanti f icat ion profi les

The three ImageQuant profiles above show the quantification profile for purified DNA samples that are untreated or treated with topoisomerase I or topoisomerase II. The profiles display the difficulty in quantifying catenanes as represented by (1) and (3) because of the large monomer peak (2) between them.

Chapter 4 Results

115

Figure 4.12 Nocodazole arrest shows persistent RCIII catenanes

Wild type cells containing RCIII (Y4259) were synchronised in G1 and then released into media containing nocodazole causing metaphase arrest. DNA replication coincides with the appearance of a distinct catenated band.

Figure 4.13 Condensin deplet ion phenotype closes matches nocodazole

arrest

Brn1 was inactivated in RCIII containing cells (Y4264) by ts allele. Cells were synchronised in G1 and then released into media at the non-permissive temperature and allowed to progress through one cell cycle before re-arrest.

Chapter 4 Results

116

Figure 4.14 Nuclear deplet ion of Brn1

Brn1 was depleted from RCIII containing cells (Y4327) by anchor away. Cells were synchronised in G1 and then released into media containing rapamycin and allowed to progress through one cell cycle before re-arrest.

Chapter 4 Results

117

At this point, we had shown that condensin is required for proper and complete

removal of topological links between sister chromatids and consequently, depletion

of condensin results in chromosome segregation defects. Therefore the next crucial

issue to address was why this occurs. As discussed in the introduction, condensin

itself is incapable of resolving topological linkages and this must be accomplished

by the enzyme topoisomerase II. The question thus remains, how does the removal

of condensin affect topoisomerase II, resulting in unresolved catenation? In the

next results chapter, we will share the results of our investigations into the

mechanics of the relationship between condensin and topoisomerase II.

Chapter 5. Results

118

Chapter 5. Results: Investigating Interactions

5.1 Do condensin and topoisomerase II directly interact?

In order to elucidate the mechanism by which condensin promotes sister

chromatid decatenation, we asked whether we could detect a direct protein

interaction between condensin and topo II. We fused the endogenous TOP2 gene,

encoding topo II, to a Pk epitope tag to facilitate immunoprecipitation and detection

(Craven et al., 1998). The condensin subunit Brn1 was fused to an HA epitope.

Brn1-HA was efficiently recovered in topo II-Pk immunoprecipitates, but not in a

control immunoprecipitate from a strain lacking the topo II Pk tag (Figure 5.1A).

This suggests that condensin interacts with topo II in yeast cell extracts.

However, doing the co-precipitation experiments by just using cell lysate

could be generating false positives. As has been shown previously in the lab

(D'Ambrosio et al., 2008b), the distribution of condensin and topoisomerase II is

very similar along the DNA. Therefore, positive results for co-precipitation could be

generated not by genuine interaction between the proteins, but instead by

condensin and topoisomerase II both binding the same fragments of DNA in close

proximity, effectively co-precipitating them via their association with the same

stretch of DNA.

To overcome this potential false positive, we used a technique that more

gently lysed the cells and allowed us to pellet the chromatin without fragmenting

the DNA. In this way we collected the supernatant fraction, which would contain no

DNA, and using this fraction only, tested for the interaction of condensin and

topoisomerase II (Figure 5.1B). In this experiment, we now performed a Brn1-HA

immunoprecipitation. However, in the HA pull-down, neither Brn1-HA nor Top2-Pk

was recovered. It seems that protein stability was compromised in this type of

extract.

To test this possibility, we returned to our original method of lysing the cells

such that DNA was present in the whole cell extract and could mediate any

possible interaction between the two proteins. However, we required an additional

control to ensure that the original interaction observed was specific, and not just a

consequence of the abundance of the two proteins. For this, we included the

additional control of tagging cohesin (specifically Scc1) with the same HA epitope

Chapter 5. Results

119

tag that Brn1 was tagged with. By performing a topo II-Pk immunoprecipitation, we

could see if the enzyme pulled down both proteins with it, or specifically just one

(Figure 5.1C). Examining the results, we could again clearly detect condensin in

the pull-down but we saw cohesin being pulled down as well. However, the single-

tagged Scc1-HA6 control strain also exhibited a faint band in the pull down,

indicating that there was some unspecific binding of cohesin by the beads. Despite

further repeats, we were unable to get rid of the background cohesin binding by the

beads, leaving us unable to make a clear decision on the nature of interaction

between condensin and topoisomerase II. At this point, it seemed clear that we

would be unable to obtain much more information from these sets of co-

immunoprecipitation experiments, so we considered how else biochemically we

could investigate their interaction.

Chapter 5. Results

120

Figure 5.1 Co-immunoprecipitat ion of condensin and topoisomerase II

(A) Co-immunoprecipitation of condensin with topo II from yeast whole cell extracts. Topo II-Pk was immunoprecipitated using an antibody directed against its Pk epitope tag. Western blots of the input and unbound fractions, as well as the immunoprecipitates from strains containing the indicated epitope tag combinations are shown. (B) HA epitope pull-down was performed on cell lysate supernatant containing no DNA. (C) The same experiment as (A) but the extra control of Scc1-HA6 was included to verify if the interaction observed between condensin and topoisomerase II was specific.

Chapter 5. Results

121

5.2 Purif ication of topoisomerase II and condensin

To really be able to determine whether and how condensin and topo II

interact, we decided to investigate whether biochemically purified condensin and

topo II interact. To do this, we first purified topo II from yeast cells overexpressing

the enzyme, following a published protocol (Worland and Wang, 1989). In Figure

5.2 we can see the steps of this effective purification technique, which yielded

enzyme preparations in which topo II was the only protein detectable by

Coomassie blue staining and was functionally active in trial decatenation and

relaxation assays.

For the purification of condensin, the protein was affinity purified from yeast

strain overexpressing the five condensin subunits (St-Pierre et al., 2009). Initially

we attempted to purify condensin using the two-step purification protocol detailed in

their paper, but had faced problems with low yields of condensin that consisted of

subunits that were not stoichiometrically correct. In collaboration with Céline

Bouchoux in the lab, we were instead able to purify condensin in a one step

procedure using an anti-HA affinity matrix that specifically bound condensin.

Chapter 5. Results

122

Figure 5.2 Topoisomerase II purif icat ion steps

Overexpression of topoisomerase II is induced through addition of galactose. After 6 hours, cells are collected, lysed and topoisomerase II is purified through a two-step process of binding and elution with cellulose phosphate and Q-sepharose.

Chapter 5. Results

123

5.3 Condensin and topoisomerase II directly interact

Having purified both protein complexes, we were now able to determine the

nature of their interaction. To do this, condensin was bound via an HA epitope-

tagged Brn1 subunit to α-HA antibody-coupled affinity beads, then purified topo II

was added (Figure 5.3). Before performing the pull-down, both preparations

underwent nuclease treatment. As a control, we added topo II to the α-HA affinity

beads in the absence of condensin. After washes, bound proteins were eluted and

analyzed. Topo II was efficiently recovered from condensin-bound, but not control,

beads. This suggests that budding yeast topo II has a direct and physical

interaction with condensin.

5.4 Condensin stimulates in vitro DNA decatenation by

topo II

Given the physical interaction between condensin and topo II, we wondered

whether condensin has a direct impact on the ability of topo II to resolve catenated

DNA. As an assay to measure the decatenation activity of topo II, we used

kinetoplast DNA (kDNA) as the substrate (Marini et al., 1980). kDNA is a

mitochondria-derived network of intercatenated DNA minicircles of approximately

2.5 kb in size. Prior to decatenation, kDNA will remain in the wells during

electrophoresis, owing to its high aggregate molecular weight. In the presence of

topo II, decatenated minicircles are released that enter the gel. Incubation with our

purified yeast topo II preparation led to efficient kDNA decatenation (Figure 5.4).

DNA decatenation by topo II is an ATP-dependent reaction, and omission of ATP

from the incubation prevented minicircle release. This serves as a control to ensure

that we are observing topo II-associated decatenation activity and that gel entry is

not due to a contaminating nuclease activity.

To study the effect of condensin on decatenation, we reduced the topo II

concentration in the reaction such that, in the absence of condensin, approximately

10% of the kDNA substrate was decatenated during the incubation. Addition of

increasing concentrations of condensin led to a marked, dose-dependent,

stimulation of kDNA decatenation. In the presence of 20 nM condensin,

Chapter 5. Results

124

decatenation was stimulated by almost 3-fold compared to the reaction lacking

condensin. Incubation of kDNA with condensin in the absence of topo II did not

result in minicircle release. This suggests that condensin stimulates kDNA

decatenation by topo II.

The reaction mix in which we observed the greatest stimulation of

decatenation by condensin contained approximately equimolar amounts of kDNA,

topo II and condensin. Considering that condensin and topo II interact, condensin

might therefore directly stimulate the catalytic activity of topo II by binding to it.

Alternatively, condensin might introduce a conformational change to the DNA

substrate that facilitates its decatenation by topo II. To differentiate between these

possibilities, we investigated whether the ability of budding yeast condensin to

enhance kDNA decatenation is restricted to topo II from the same species, or

whether it extends to type 2 topoisomerases from other organisms. We used

commercial preparations of human topo IIα and E. coli topo IV in this comparison

(Figure 5.5). Topoisomerase concentrations were again titrated such that in the

absence of condensin approximately 10% of the kDNA substrate was resolved.

Addition of condensin in each case led to a reproducible stimulation of the kDNA

decatenation reaction. Human topo IIα shows 49% sequence identity and can

functionally complement the essential in vivo function of yeast topo II (Jensen et al.,

1996). A protein interaction between the two proteins could therefore be conserved

between the species. This is less likely the case for E. coli topo IV, even though it’s

parE subunit shows 24% identity and 43% similarity to the N-terminal half of

budding yeast topo II. Condensin might thus promote decatenation both by

establishing a DNA substrate geometry that facilitates decatenation as well as

recruit or activate topo II to assist the decatenation reaction.

Chapter 5. Results

125

Figure 5.3 Condensin interacts direct ly with purif ied topoisomerase II

Purified topo II binds to condensin that is retained on an antibody affinity matrix. Purification and interaction analysis was performed as described in Materials and Methods, the input and eluates were analyzed by SDS-PAGE and proteins visualised by Coomassie blue staining.

Chapter 5. Results

126

Figure 5.4 Condensin promotes decatenation of kinetoplasts

100 ng kDNA (corresponding to a concentration of 3 nM minicircles) were incubated with the indicated concentrations of purified yeast topo II and condensin. Reactions lacking topo II or lacking ATP were included as controls. Reaction products were resolved by agarose gel electrophoresis and visualised by ethidium bromide staining.

Chapter 5. Results

127

Figure 5.5 Condensin st imulates dif ferent topoisomerase II enzymes

Figure 5.5 as per Figure 5.4, but including reactions using E. coli topo IV and H. sapiens topo IIα. Minicircle release was quantified in at least 3 independent experiments with each topoisomerase. The mean and standard deviations are shown.

Chapter 6. Results

128

Chapter 6. Results: a-factor Synthesis and

Characterization

6.1 Synthesis of a-factor

As was introduced in chapter 1.7, the anchor away strains we used were

mostly mating type α , which meant that to synchronise the cells in G1 the mating

pheromone a-factor had to be used. However, while α-factor is readily produced by

peptide synthesis laboratories in many research institutes as well as being

commercially available, a-factor is not. This is because α-factor lacks any peptide

modifications while a-factor has both farnesylation and carboxyl methylation, both

of which are important for the biological potency of a-factor (Anderegg et al., 1988)

(Marcus et al., 1991). This makes α-factor readily accessible by automated solid-

phase peptide synthesis and much easier to manufacture. While over the years

many a-factor synthesis strategies have been published, they typically involve

liquid-phase reactions that are a hallmark of expert organic chemistry laboratories

and thus out of reach for many users who do not have access to such expertise

(Xue et al., 1989) (Sherrill et al., 1995) (Mullen et al., 2011).

In collaboration with the Peptide Synthesis laboratory at our institute, a

novel strategy for the synthesis of a-factor was developed. The process is based

on solid-phase peptide synthesis, followed by two simple steps in solution using

widely available reagents and apparatus, thus making a-factor accessible to

laboratories with standard peptide synthesis facilities (O'Reilly et al., 2012). Our

contribution was to demonstrate the successful use of our synthetic a-factor to

synchronise cell cycle progression of cells of the α mating type.

6.2 Use of a-factor to synchronise cells of α mating type

Initially we tried to synchronise haploid cells of the α mating type, using a-

factor concentrations and conditions similar to what is used for α-factor

synchronization of a mating type cells. Adding a-factor at a concentration of 0.5

µg/ml to asynchronously growing cells in early exponential phase led to efficient

cell cycle arrest in G1. However, upon filtration and washout of the a-factor, these

Chapter 6. Results

129

cells resumed cell cycle progression only later and very inefficiently. We therefore

reduced the a-factor concentration through a series of optimisation experiments.

Eventually we identified that addition of a total concentration of 0.04 µg/ml a-factor,

in two halves at the beginning and after 1 h of arrest, efficiently arrested cells in G1.

Washout of the a-factor and resuspension in fresh pheromone-free medium now

resulted in synchronous release from the arrest (Figure 6.1). We followed cell cycle

progression by FACS analysis of DNA content, as well as by scoring the budding

index and the fraction of binucleate cells. 60 minutes after release, when most cells

showed a new bud, we added a-factor back to the culture to re-arrest cells after

completion of mitosis in the following G1. This analysis showed that arrest of α cells

with a-factor and release by filtration gives rise to a cell population that transverses

the cell cycle with very good synchrony.

The a-factor concentration we used in this protocol is 10-fold lower than the

concentration of 2-factor that we routinely use to synchronise a mating type cells. It

is 100-fold lower than recommended α-factor concentrations in some

synchronization protocols (Breeden, 1997). When we tried to use α-factor at a low

concentration to arrest a mating type cells, we found that 0.04 µg/ml α-factor was

not sufficient to impose a stable cell cycle arrest. We do not know the reason for

the greater specific biological activity of a-factor as compared to α-factor. Its

hydrophobic nature and farnesylation might enhance its affinity for lipid membranes

and thereby increase its local concentration close to the target cell a-factor receptor.

In addition, the absence of a known a-factor protease, equivalent to the Bar1 α-

factor protease, may mean that a-factor is more stable in the culture medium or in

the vicinity of the receptor.

Chapter 6. Results

130

Figure 6.1 Use of a-factor to synchronise cel l cycle progression of α cel ls

0.02 µg/ml a-factor was added to a culture of exponentially growing budding yeast cells of α mating type. After 60 minutes, a second 0.02 µg/ml dose of the pheromone was added. After 120 minutes, all cells were arrested in G1 (time point 0). The culture was then filtered and the cells were washed and resuspended in fresh medium lacking a-factor. The cells now passed through a synchronous cell cycle and a-factor was added to the culture again after 60 minutes to impose re-arrest in the following G1. Cell cycle progression was monitored by FACS analysis of DNA content and microscopic analysis of the budding index and binucleate cells.

Chapter 6. Results

131

6.3 Litt le shmoo formation during a-factor-induced G1

arrest

When we microscopically followed the a-factor response during cell

synchronization, we noticed that even though cells arrested unbudded, they

showed hardly any signs of shmoo formation. This was unlike α-factor-treated a

mating type cells, where G1 arrest is typically accompanied by noticeable shmoo

formation (Figure 6.2). In our cultures, α cells treated with synthetic a-factor lost

their oval shape and took on a more rounded, sometimes lemon-shaped,

appearance. The cell volume increased over time in the arrest, but directed shmoo

formation was barely observed. This was even the case at higher pheromone

concentrations, similar to those that elicit efficient shmoo formation of α-factor-

treated a mating type cells (Figure 6.2). While it has been previously noticed that α

cell shmoos differ from the typical a cell projections (Wilkinson and Pringle, 1974)

(Betz et al., 1977), our observations suggest that as yet poorly understood

differences exist in the way the opposite mating pheromones act on yeast cells, or

in the way that cells of the opposite mating types react to pheromone exposure.

Chapter 6. Results

132

Figure 6.2 Comparison of pheromone-induced shmoo formation of a and α

cel ls

Haploid cells of either a or α mating type were grown in early exponential phase. Mating pheromones α- and a-factor, respectively, were added at the indicated concentrations. Cells were photographed before and 2.5 hours after pheromone addition, using a Zeiss Axioplan 2 microscope equipped with differential interference contrast optics.

Chapter 7. Discussion

133


7.1 Catenation can be observed in minichromosomes

In our experiments, we have shown the utility of our catenation assay in

observing and tracking catenanes for a range of minichromosome sizes. Despite

previous reports that replication-dependent catenation of smaller minichromosomes

was too transient to be detectable in wild type cells (Baxter et al., 2011), we could

observe catenation with all our minichromosome substrates. By careful optimisation

of our DNA purification process and electrophoresis conditions, we established

parameters that enabled us to clearly track catenane behaviour through the cell

cycle and our purified minichromosomes show distinct patterns of replication-

dependent catenation that resolve with a specific timing.

The significance of our assay results was further supported by the dual

approaches we took towards protein inactivation. By being able to inactivate

proteins by both ts allele disruption and anchor away-mediated nuclear depletion,

we were able to verify that the resultant phenotypes were indeed a specific result of

the target protein’s inactivation.

7.2 Cohesin protects catenation

When cohesin was inactivated through either use of a ts Scc1 allele or

nuclear depletion via anchor away, we saw a significant reduction in the abundance

of catenanes. During S-phase, when we normally see the greatest proportion of

catenanes, less than 20% of minichromosomes were detected as catenated. Thus

it appears that cohesin plays a substantial role in the protection of catenation.

There are differing mechanisms by which cohesin could exert this influence; an

‘indirect’ mechanism whereby cohesin might retard topo II-driven decatenation

hence maintaining topological linkages, or ‘directly’ through the entrapment of sister

DNAs within the cohesin tripartite ring. In the former mechanism, cohesin could

retard decatenation by topo II through some sort of interaction. In our co-

immunoprecipitation studies in chapter 5.1, we used cohesin as a control and saw

an unexpected interaction between Scc1 and Top2, which suggests that some


134

direct interaction between the two proteins is at least feasible. Another possibility is

that the cohesin complex blocks topo II’s access to sites of intertwining, inhibiting

decatenation. As the cohesin tripartite ring physically entraps the sister DNAs, it is

possible that the sites of entrapment coincide with where the sister DNAs are

physically closest, or catenated. The presence of cohesin could therefore block

access until the time it is cleaved by separase.

However, if we take into account the effect of other mitotic components on

catenation, it seems that cohesin may play a more indirect role. If we compare

catenane levels between cells arrested in metaphase either by nocodazole or by

inactivation of Cdc20, we can see that there is a marked decrease in catenanes

when the spindle is present. In both situations cohesin is still present and

uncleaved, yet the presence of the mitotic spindle seems to promote decatenation,

implying that the cohesin complex isn’t blocking topo II. Thus, cohesin could be

playing a more direct role in maintaining catenation. We hypothesize that while

topo II is able to access and act upon the sister DNAs, prior to cohesin cleavage,

those sister DNAs will remain physically intimate owing to their entrapment by the

cohesin ring. Therefore, the reintroduction of topological linkages owing to the bi-

directionality of topo II is possible. As long as cohesin remains uncleaved, an

equilibrium of decatenation and recatenation will be established between sister

DNAs. The influence of other factors, such as the tension exerted by the mitotic

spindle or the cleavage of cohesin allowing sister DNAs to move away from each

other, will alter the balance of that equilibrium. A recent study has come to a similar

conclusion on cohesin’s catenane-protection properties (Farcas et al., 2011).

7.3 Anaphase bridges result from persistent sister

chromatid catenanes

Chromosome segregation failure and anaphase bridges are a hallmark

phenotype of cells with compromised condensin function, yet prior to our study, the

reason underlying this failure has remained undetermined. We now provide

evidence that, at least in the case of circular minichromosomes in budding yeast,

condensin is required to promote complete resolution of catenanes that are

retained between sister chromatids following their replication. Persistent sister


135

chromatid catenanes have previously been proposed as a source for anaphase

bridges in a condensin mutant (Bhat et al., 1996), though it is to our knowledge the

first time that these catenanes have been directly observed. An important

prediction from a model where sister chromatid catenation prevents chromosome

segregation is that ectopic decatenation should resolve the anaphase bridges in

the absence of condensin. This has been achieved in case of the budding yeast

rDNA locus where anaphase bridges in a condensin mutant were resolved by

ectopic expression of a viral topo II enzyme. Though again the catenations status

of the locus could not be observed at the time (D'Ambrosio et al., 2008a). Taken

together with the direct visualization of persistent catenanes between circular

chromosomes that we now report, we can suggest the chromosome bridges that

arise when condensin function is compromised are indeed due to persistent DNA

catenation.

7.4 Condensin promotes decatenation by topoisomerase

II

An important area for future research is to investigate whether catenation

between linear chromosomes is regulated in the same way as we have observed

here in the case of circular substrates. Catenation, in theory, should not exist

between linear pieces of DNA, as any intertwining could easily slide off the DNA

ends. This is a reason why catenation of linear chromosomes cannot be studied

after DNA isolation by gel electrophoresis. In vivo, however, chromosome

movement is constrained such that catenation between linear sister chromatids is

maintained and has the ability to restrain chromosome segregation during

anaphase (Holm et al., 1985, Uemura et al., 1987b). Whether, and how far,

topological links between sister chromatids might be able to translocate along DNA

strands is not known. Helical tension along individual sister strands is constrained

to regions of approximately 100 kb in budding yeast (Joshi et al., 2010). It seems

likely that sister chromatid catenation is similarly constrained. Topologically isolated

domains along chromosomes are probably the consequence of protein-mediated

DNA interactions, as has been demonstrated at least in the case of mitotic

chromosomes (Kawamura et al., 2010). How topological interlinks between linear


136

chromosomes arise, where and for how long they are maintained, and how they

are eventually resolved are important question. To address these will likely require

the topological isolation of sections of linear chromosomes by excision in a circular

form to facilitate their analysis.

7.5 Decatenation occurs in steps

While studying the sister chromatid catenation status of circular

chromosomes during the cell cycle, we found that most catenanes are rapidly

resolved by topo II soon after DNA replication. When topo II was active, at most

half of the chromosomes were seen catenated in our synchronised cell population

at any one time. Only 10-20% catenanes persisted until mitosis, or in the absence

of cohesin when proteinaceous sister links were removed. This fraction is similar to

the fraction of catenanes that persisted throughout mitosis and into G1 of the next

cell cycle in cells depleted of condensin. This raises the question whether there is a

distinction between catenanes that are readily resolved and those that are

decatenated later in the cell cycle and require condensin for their resolution? We

can see two possible scenarios. In the first, all replication products are left

catenated in the same way, possibly by a small number of interlinks per replication

termination site. Topo II will act on these based on chance encounters sister DNA

strands. Based on its mode of DNA binding, topo II favors decatenation over

catenation at such crossing points (Vologodskii et al., 2001, Dong and Berger,

2007). Resolution is limited in this scenario by Brownian movement of sister

chromatids, constrained by cohesin-dependent cohesion. Condensin’s role would

be to accelerate resolution by stimulating topo II activity and promoting a substrate

geometry conducive to decatenation. In an alternative scenario, topological

interlinks between sister chromatids might exist in different forms. Depending on

the reactions involved in replication termination and constraints imposed by the

chromosome environment the catenanes produced might differ in the number of

interlinks or additional topological features, e.g. knots. Simple catenanes could be

good substrates for rapid decatenation, while condensin would aid the resolution of

more complex topologies. In S. cerevisiae, the rDNA shows the greatest resolution

problems in condensin mutants (D'Ambrosio et al., 2008a). This area of DNA is


137

highly transcribed, even during mitosis, possibly generating topologically complex

catenanes that benefit more from condensin activity. Condensin binding to DNA

introduces positive-handed writhe, which in the presence of topo II allows knot

formation (Kimura et al., 1999). Equally, by stabilizing transient DNA conformations

that rarely form spontaneously, condensin might facilitate resolution of complex

structures. In support of this possibility, the fraction of catenanes that require

condensin for resolution appear to be refractory to decatenation in its absence and

persist for extended time periods. Such structures might also pose the greatest

danger to chromosome segregation.

7.6 Condensin directly interacts with topoisomerase II

How does condensin promote sister chromatid decatenation? As discussed

above condensin might act at the level of the DNA substrate to facilitate catenane

resolution, maybe in particular those of a complex topological nature. Our

observation that budding yeast condensin facilitates the in vitro decatenation of

kDNA circles not only by budding yeast topo II but also by topoisomerases of

human and bacterial origin is consistent with such an activity. In addition to

affecting DNA conformation, we also observed a direct protein interaction between

budding yeast condensin and topo II. Such an interaction could directly stimulate

the catalytic activity of topo II as has been recently observed in the bacterial system

(Li et al., 2010, Hayama and Marians, 2010). Yet another way how condensin’s

interaction with topo II could facilitate sister chromatid decatenation is by

recruitment of topo II to its sites of action. Condensin and topo II show significant

overlap in their chromosomal association pattern (D'Ambrosio et al., 2008b),

consistent with a role in topo II recruitment.

7.7 Future perspectives

Our catenation assay has proved to be a useful and flexible tool in our

investigation, having produced an array of interesting results. Coupled with our in

vitro biochemical experiments using purified proteins, we have been able to make

some novel insights into condensin’s and topoisomerase II’s role in chromosome


138

resolution. However, looking to the future we believe that the next logical step

would be to move on from using minichromosomes as models and to develop

techniques to study catenation on the native chromosomes themselves. We believe

that we have reached the limit of what can be accurately observed using

minichromosomes. In our experiments, as minichromosome size increased so did

linearization of the substrate, most likely owing to an increased vulnerability to

pipette-induced shearing. We found that by minimizing the steps in our purification

protocol that involved pipette use, as well as implementing other purification

optimizations, we could minimise the amount linearised. However, it is likely that

our assay has pushed the limits of what is observable with large sized

minichromosomes. RCIII, being over 60 kb in size, already presented many

difficulties in resolution and was very susceptible to damage. While we found that

pulse field gel electrophoresis did not yield satisfactory resolution for our assay, it is

likely that any larger substrates would have to be resolved via this technique.

However, we feel that the next logical step for future investigations would be the

formation of a technique that could manipulate the native chromosomes

themselves, a direction that some labs have already tentatively begun moving in

(Farcas et al., 2011). It is feasible that native chromosomes could be engineered

such that sections could be cut out and ligated in vivo to create circular DNAs that

are easily assayable by adapting our current protocols, perhaps through adaption

of the Cre/lox recombinase system commonly used in mammalian and plant cells

(Figure 7.1). With the availability of condensin binding maps, replication origins and

knowledge of areas of potential catenation, interesting sites could be marked for

excision and study finally allowing for the direct observation of endogenous

chromosomes.


139

Figure 7.1 Schematic displaying possible excision and re-l igation of

endogenous chromosome DNA to form de novo r ing chromosomes in vivo

In the figure two sister DNAs are topologically linked and cut sites (red boxes) flank the site of catenation. Upon excision at these cut sites and consequent ligation, topologically linked circular DNAs are created.


140

We are still very naïve about how cells deal with the immense topological

challenge of replicating millimeter-long chromosomes in micrometer-sized nuclei

and ensuring the smooth resolution of DNA interlinks during the segregation of

sister chromatids during mitosis. Cell division underpins the propagation of all life

on earth and unraveling this fundamental process is both fascinating and essential.

Chromosome resolution and segregation lies at the heart of human growth,

reproduction and regeneration and the danger of persisting anaphase bridges to

genome stability makes this an important area of further study.

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